Tissue scaffolds derived from forestomach extracellular matrix

ABSTRACT

The present invention pertains to the development of Extracellular Matrix (ECM) scaffolds derived from the forestomach of a ruminant. Such scaffolds are useful in many clinical and therapeutic applications, including wound repair, tissue regeneration, and breast reconstruction. In addition, the present invention features methods of isolating ECM scaffolds from mammalian organs, including but not limited to the ruminant forestomach. The invention further features laminated ECM scaffolds containing a polymer positioned between individual ECM sheets. The polymer may optionally contain bioactive molecules to enhance the functionality of the scaffold.

RELATED APPLICATIONS

This application claims priority to U.S. Provisional Application No.61/137,367, filed on Jul. 30, 2008, and to U.S. Provisional ApplicationNo. 61/172,671, filed on Apr. 24, 2009. The entire contents of theforegoing applications are incorporated herein by reference.

BACKGROUND OF THE INVENTION

Extracellular matrix (ECM) has an important role in providing theoptimal chemical and structural environment for tissue growth andregeneration. ECM scaffolds used for tissue regeneration aretraditionally prepared from decellularised human and animal dermisisolated from various organs, and from a variety of animal submucosaland basement membrane sources. These scaffolds promote tissueregeneration and are well-tolerated immunologically. Common submucosaltissue graft compositions are derived from the small intestine, theurinary bladder, and the simple glandular stomach (see, for example,U.S. Pat. No. 4,902,508, U.S. Pat. No. 5,554,389, and U.S. Pat. No.6,099,567, the entire contents of which are incorporated herein byreference).

Despite advances in the production and use of ECM scaffolds, the idealscaffold composition has not been identified. The ideal scaffold isnon-allergenic, non-carcinogenic, mechanically stable under continuousstress, adequately porous to allow capillarisation, able to encourageand direct appropriate cellular and vascular in-growth, of similarcompliance to that of the host tissue, resistant to infection,non-thrombogenic, inexpensive, and able to become a fully functionalanalog of the original tissue. An ECM scaffold possessing theseproperties would be useful in a variety of clinical applications,including wound repair and soft tissue regeneration.

SUMMARY OF THE INVENTION

The present invention provides extracellular matrix (ECM) scaffoldsderived from the forestomach of a ruminant, also referred to herein as“Forestomach Matrix” (FM) scaffolds. The FM scaffolds of the inventionprovide a number of advantages over prior tissue scaffolds, and areuseful in a variety of clinical and therapeutic applications, includingwound repair and tissue regeneration. In addition, the present inventionprovides improved methods of producing ECM scaffolds from mammalianorgans, including but not limited to the ruminant forestomach. Inparticular embodiments, FM scaffolds of the invention may be derivedfrom a ruminant belonging to the genus Capra, Bos, Cervus or Ovis, e.g.,Capra aegagrus hircus, Bos taurus, or Ovis aries.

Accordingly, in a first aspect, the invention features a tissue scaffold(an FM scaffold) comprising the ECM of the propria-submucosa of theforestomach of a ruminant. In a particular embodiment, thepropria-submucosa is from the rumen, the reticulum, or the omasum of theforestomach. These tissue scaffolds typically have a contoured luminalsurface. The ECM tissue scaffolds of the invention may additionallycontain decellularised tissue, including portions of the epithelium,basement membrane, or tunica muscularis, and combinations thereof. Thetissue scaffolds may also comprise one or more fibrillar proteins,including but not limited to collagen I, collagen III, or elastin, andcombinations thereof. In other embodiments, the tissue scaffolds cancomprise one or more growth factors, including but not limited to FGF-2,TGFb1, TGFb2, or VEGF, and combinations thereof. In still otherembodiments, the tissue scaffolds can comprise one or moreglycosaminoglycans, including but not limited to hyaluronic acid andheparan sulfate, and combinations thereof. In another embodiment, thetissue scaffolds can comprise one or more adhesive proteins, includingbut not limited to fibronectin, collagen IV, or laminin, andcombinations thereof.

FM scaffolds of the present invention can be formatted in a variety ofmanners, such as a single sheet, or as a laminated sheet containingmultiple sheets of FM. In certain embodiments, the FM scaffolds comprise2-15 laminated sheets. Such laminated sheets can be held together bystitches or sutures. Alternatively, the laminated sheets can be heldtogether by a polymer positioned in between one or more sheets. In oneembodiment, the laminated sheets are attached by stitches or sutures andadditionally contain a polymer positioned between one or more sheets. Inanother embodiment, the laminated tissue scaffold comprises a polymerpositioned between each of the laminated sheets. The polymer may beinterspersed between the sheets, or may be evenly distributed as apolymer layer. Any suitable polymer may be used in the FM scaffolds ofthe invention, including but not limited to collagen, chitosan,alginate, polyvinyl alcohol, carboxymethyl cellulose, hydroxypropylcellulose, and combinations thereof.

In a particular embodiment, the polymer further comprises a bioactivemolecule, for example, a small molecule or a peptide. The bioactivemolecule may be non-covalently incorporated into the polymer, forexample, as a suspension, encapsulated as particles, microparticles, orcolloids, or as a mixture thereof. The bioactive molecule may also becovalently incorporated into the polymer, using any suitable chemistryfor attachment of the bioactive molecule to the polymer. The bioactivemolecule can be any therapeutically desirable molecule, such as a growthfactor, an anti-microbial, an analgesic, a hemostatic, a pro-angiogenicagent, or an anti-angiogenic agent. In exemplary embodiments, thepolymer comprises one or more of FGF2, NGF, doxycycline, amoxicillin,and poly-L-lysine.

In another particular embodiment, the FM scaffolds have a width of atleast 10 cm. For example, the scaffold can have a width of at least 10cm and a length of at least 10 cm. Accordingly, certain FM scaffolds canhave a surface area of more than 100 cm², e.g., 400 cm². In oneembodiment the single or laminated sheets of an FM scaffold areperforated. In another embodiment the FM is fluidized, or micronized.

FM scaffolds of the invention generally have a biaxial strength greaterthan scaffolds obtained from other gastrointestinal or urogenitalsources. Accordingly, in a particular embodiment, the FM scaffolds havean average biaxial strength of at least 80 N or more.

Tissue scaffolds of the invention can be used in multiple applications,including but not limited to covering a tissue deficit and reinforcingsoft tissue. In a particular embodiment, the tissue deficit or the softtissue has width of at least 10 cm. In another embodiment, the tissuedeficit or the soft tissue has a width of at least 10 cm and a length ofat least 10 cm. In still another embodiment, the tissue deficit or thesoft tissue has a surface area of at least 100 cm².

Accordingly, in another aspect, the invention features a method forinducing repair of a damaged tissue, comprising contacting the damagedtissue with an FM scaffold of the invention, e.g., one that comprisesthe ECM of the propria-submucosa of a ruminant. The invention furtherfeatures a method for stimulating soft tissue regeneration, comprisingcontacting the soft tissue with an FM scaffold of the invention.

When an FM scaffold is placed in contact with a tissue, the FM scaffoldcan increase proliferation of cells located near the scaffold. Inaddition, the FM scaffold can promote vascularization within a tissue towhich it adheres. Accordingly, in another aspect, the invention providesa method of stimulating proliferation of cells in a tissue, comprisingcontacting the tissue with an FM scaffold such that cell proliferationis stimulated. The invention further provides a method of inducingvascularization of a tissue, comprising contacting the tissue with an FMscaffold such that vascularization occurs within the tissue.

In one aspect, the invention features an implantable tissue scaffolddevice for supporting breast tissue within a patient, wherein the devicecomprises extracellular matrix of the propria-submucosa of theforestomach of a ruminant. The breast tissue may comprise a breastprosthesis, i.e., a breast implant. The tissue scaffold device may beformatted as a laminated sheet comprising two or more layers ofextracellular matrix. In a particular embodiment, the laminated sheetcomprises 2-15 layers of extracellular matrix.

The tissue scaffold device may be flat, or it may have a concavity. Inone embodiment, the layers of extracellular matrix of the device may besecured together by stitches or sutures. The extracellular matrix may beperforated, or it may be unperforated. In some embodiments, the devicehas a crescent shape. In other embodiments, the device has an ellipticalshape.

In another aspect, the invention provides a method of supporting breasttissue within a patient, comprising positioning the foregoing tissuescaffold device within the patient in a supporting position relative tothe breast tissue. In one embodiment, the breast tissue comprises abreast prosthesis. In another embodiment, the breast tissue comprisesnative tissue. In a particular embodiment, positioning the tissuescaffold comprises covering the lower and lateral sections of the breasttissue.

In another aspect, the invention provides a tissue scaffold comprisingtwo or more sheets of extracellular matrix, laminated by a polymerpositioned between the sheets. The scaffold may comprise extracellularmatrix of the submucosa of a tissue selected from the group consistingof small intestine, stomach, bladder, pericardium, and dermis. In aparticular embodiment, the extracellular matrix comprises collagen. Thepolymer may comprise collagen, chitosan, alginate, polyvinyl alcohol,carboxymethyl cellulose, hydroxypropyl cellulose, and combinationsthereof.

In a particular embodiment of the foregoing aspect, the polymer furthercomprises a bioactive molecule. The bioactive molecule may benon-covalently or covalently linked to the polymer. In one embodiment,the bioactive molecule may be a small molecule or a polypeptide, e.g., agrowth factor, an anti-microbial, an analgesic, a hemostatic, apro-angiogenic agent, an anti-angiogenic agent, or combinations thereof.Exemplary bioactive molecules include FGF2, NGF, doxycycline,poly-L-lysine, and combinations thereof.

In yet another aspect, the present invention provides methods ofgenerating ECM tissue scaffolds by separating and/or decellularising thelayers within all or a portion of a tissue. The methods involve creatinga transmural osmotic flow between two sides of the tissue, such that thelayers within all or a portion of the tissue are separated and/ordecellularised. The transmural osmotic flow can be directed from theluminal to the abluminal side of all or a portion of the tissue, or fromthe abluminal to the luminal side of all or a portion of the tissue.This can be achieved, for example, by separating the tissue between ahypertonic and a hypotonic solution, such that the transmural osmoticflow is directed from the hypotonic solution to the hypertonic solution.The method can further involve removing all or part of a tissue layerincluding epithelium, basement membrane, or tunica muscularis, andcombinations thereof.

In a particular embodiment, the method of the invention involvesencapsulating a first solution within an organ or tissue (or a portionthereof), and immersing the organ or tissue (or portion thereof) in asecond solution which is hypertonic to the first solution. This methodcan further involve removing the organ or tissue from the secondsolution, and immersing the organ or tissue, or portion thereof, in athird solution which is also hypertonic to the first solution, in orderto, for example, further decellularise the tissue.

In an alternative embodiment, the method comprises encapsulating a firstsolution within an organ or a tissue (or portion thereof), and immersingthe organ or tissue (or portion thereof) in a second solution which ishypotonic to the first solution, optionally followed by immersing theorgan or tissue, or portion thereof, in a third solution which is alsohypotonic to the first solution.

The hypertonic and hypotonic solutions can include, for example, waterand optionally at least one buffer, detergent or salt. The hypertonicsolution contains a higher concentration of solute than the hypotonicsolution. In a particular embodiment, the hypertonic solution comprises4 M NaCl, and the hypotonic solution comprises 0.028% Triton X-200 and0.1% EDTA. In another particular embodiment, the hypotonic solutioncomprises 0.1% SDS. In still another embodiment, the hypotonic solutioncomprises 0.028% Triton X-200, 0.1% EDTA, and 0.1% SDS.

The methods of the invention can be performed at low temperatures of,for example, 4° C. or less (e.g., between about 2° C. and about 4° C.).The methods of the invention can alternatively be performed at or nearroom temperature (e.g., between about 18° C. and about 24° C. Themethods also allow tissue layers to be separated and decellularised in ashorter period of time than is possible using other methods. Inparticular embodiments, the tissue layers are separated anddecellularised in 36 hours or less, more preferably in 24 hours or less.In other embodiments, the tissue layers are separated and decellularisedin 6 hours or less (e.g., in 5 hours or less, 4 hours or less, or 3hours or less).

The methods of the invention can be employed with any suitable tissuesource or organ. In a particular embodiment, the tissue comprises akeratinized stratified squamous epithelium. In other particularembodiments, the tissue is derived from the forestomach of a ruminant,e.g., an animal belonging to the genus Capra, Bos, Cervus and Ovis. Instill other embodiments, the tissue is derived from the rumen, thereticulum, or the omasum of the forestomach. Such tissues can optionallybe distended to increase the transmural osmotic flow across the tissuelayers, further facilitating separation and decellularisation.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1 is a schematic cross-section of (A) the forestomach wall, and (B)the glandular stomach wall, both in an unprocessed state and after STOFprocessing.

FIG. 2 illustrates two exemplary shapes of FM scaffolds useful in breastaugmentation, reconstruction, or mastopexy. (A) depicts a crescentshaped scaffold. (B) depicts an elliptical shaped scaffold.

FIG. 3 illustrates one embodiment of a method of processing an organ byestablishing a transmural osmotic flow across the organ.

FIG. 4 depicts an exemplary embodiment of tissue processing using STOFto produce a decellularised FM scaffold with a fractured basementmembrane.

FIG. 5 shows the total nucleic acid content of tissue before and afterthe STOF process.

FIG. 6 shows the linear relationship between surface area of forestomachtissue and increases in total forestomach volume after the STOF process.

FIG. 7 shows the change in tissue weight as a result of transmuralosmotic flow. Fluid passing through and into the tissue resulted inweight gain as tissue hydration increased.

FIG. 8 shows a Western blot detection of laminin in STOF solutions atcompletion of the STOF process.

FIG. 9 presents results of the Ball Burst test, showing the relativestrength of FM.

FIG. 10 graphically depicts the ball burst strength of multi-ply ovineFM products. The biaxial strength of single or multi-ply ovine FMproducts was determined using a Ball Burst Test according to ASTM D3797-89 “Standard Test Method for Bursting Strength of Knitted Goods,Constant-Rate-of-Traverse (CRT) Ball-burst Test”, using an Instron 5800series electromechanical tester, fitted with a ball-burst compressioncage, whereby a 25.4 mm stainless steel ball was pushed against the testmaterial at a feed-rate of 305+/−13 mm/min, until failure. A 1 kN loadcell was used to record maximum compression load at failure (N). Errorbars represent the standard error of at least five samples.

FIG. 11 depicts a comparison of normalized maximum compression load for4-ply ovine FM and commercially available implant products. Error barsrepresent the standard error of at least five samples, or from publisheddata.

FIG. 12 presents results of the Uniaxial test, showing the relativestrength of FM.

FIG. 13 presents a comparison of the strength of ovine FM single andmulti-ply products. (A) Maximum load at failure (N); (B) Maximumtangential stiffness (N/mm); (C) Maximum elongation (mm); (D) Modulus ofelasticity (Young's) (GPa); (E) Yield Stress (MPa); and (F) thickness.Maximum load at failure of single and multi-ply products was determinedusing an Instron 5800 series electromechanical tester. Various materialswere cut to dog-bone shaped samples with a 0.6 cm width. Samples wereclamped with a gauge length of 7.5 cm. and elongated at a rate of 25.4mm/min until failure. Load (N) was recorded using a 500 N load cell.Stiffness was calculated from the slope of the load (N) versuselongation (mm) curve. The load versus elongation curve was transformedto a stress (N/m²) versus strain curve, using a cross-sectional area ofcalculated from the thickness of the product. The slope of this lattercurve was used to calculate the modulus of elasticity, or Young'smodulus (GPa). Error bars represent the standard error of at least fivesamples.

FIG. 14 presents a comparison of the yield stress of 1- and 2-ply ovineFM products with commercial dural repair products. Errors representstandard errors from at least five samples, or from published data.

FIG. 15 presents a comparison of the suture retention strength ofmulti-ply ovine FM products. Samples of multi-ply ovine FM products weretested for suture retention according to ANSI/AAMI VP20-1994 Guidelinesfor Cardiovascular Implants Vascular Prostheses Measured in Newton's.Sutures were made in 4 cm×2.5 cm samples, using suture with a bite-depthof 2 mm. Load at failure was recorded using a Instron 5800 serieselectromechanical tester, fitted with a 100 N load cell using an advancerate of 100 mm/min. Load at failure was defined as a 90% reduction inthe observed load. The free end of the sample was held in a 25 mm vicegrip, while the suture was attached to the opposing clamp via astainless hook. Error bars represent the standard error of at least sixsamples.

FIG. 16 presents a comparison of the normalized suture retentionstrength of ovine FM products and dural repair products. Errorsrepresent standard errors from five independent samples, or frompublished data.

FIG. 17 presents a comparison of the normalized suture retentionstrength of 4-ply ovine FM product and commercially available implantmatrices. Errors represent standard errors from at least five samples,or from published data. No error reported for Surgisis™. Alloderm™ andStrattice™ tested with a bite depth of 10 mm, 4-ply ovine FM andSurgisis™ tested with a bite depth of 2 mm.

FIG. 18 presents an example of the layout of full-thickness excisionalwounds made to the back of a pig in a porcine wound-healing study.

FIG. 19 presents the persistence of ECM scaffolds in tissue biopsiestaken during the course of the wound healing study.

FIG. 20 graphically depicts quantification of cell proliferation duringwound healing. Wounded tissue was treated with ovine FM (1-ply, 2-ply),SIS, or was untreated, and the total number of Ki67-positive cells inthree 40× frames taken from the epithelial layer and three 40× framestaken from the regenerating dermal layer were counted using IHC anddigital methods. **P<0.01 significance relative to untreated controlusing one-way ANOVA.

FIG. 21 graphically depicts a quantification of blood vessels in woundedtissue treated with ovine FM (1-ply and 2-ply), SIS, or wounded tissuethat was untreated. (A) depicts the average total number of bloodvessels counted per frame, analyzed for each tissue biopsy. Error barsrepresent standard errors from the 20 biopsies analyzed for eachtreatment group at the time points indicated. **P<0.01 and *P<0.05significance relative to untreated control using one-way ANOVA. (B)depicts the number of blood vessels (small, medium and large) as aproportion of total observed blood vessels for each of the treatments,at the time points indicated. (C) depicts the average number of smallblood vessels (300-500 μm²), counted per frame. Counts were averagedover all frames analyzed from the five animals under study. Error barsrepresent standard errors from 20 analyzed frames for each of thetreatment groups, at the time points indicated. (D) depicts the averagenumber of medium blood vessels (500-1500 dm²), counted per frame. Countswere averaged over all frames analyzed from the five animals understudy. Error bars represent standard errors from 20 analyzed frames foreach of the treatment groups at the time points indicated. (E) depictsthe average number of large blood vessels, (>1500 μm²) counted perframe. Counts were averaged over all frames analyzed from the fiveanimals under study. Error bars represent standard errors from 20analyzed frames for each of the treatment groups, at the time pointsindicated.

DETAILED DESCRIPTION OF THE INVENTION

The present invention relates to the development of Extracellular Matrix(ECM) scaffolds derived from the propria-submucosa of the forestomach ofa ruminant, referred to herein as ‘Forestomach Matrix’ (FM). FMscaffolds possess distinct characteristics which differ from ECMscaffolds derived from other organs, including the glandular stomach.These characteristics make FM scaffolds particularly well-suited forclinical applications involving tissue regeneration and repair. Thepresent invention further relates to improved methods for generating ECMscaffolds from mammalian organs, including but not limited to theforestomach.

I. DEFINITIONS

So that the invention may be more readily understood, certain terms arefirst defined.

The term “Forestomach Matrix” (abbreviated FM), as used herein, refersto an ECM scaffold containing the propria-submucosa of the forestomachof a ruminant.

The term “propria-submucosa,” as used herein, refers to the tissuestructure formed by the blending of the lamina propria and submucosa inthe ruminant forestomach.

The term “lamina propria,” as used herein, refers to the luminal portionof the propria-submucosa, which includes a dense layer of extracellularmatrix.

The term “ruminant,” as used herein, refers to a mammal having a stomachwith four chambers. These include a forestomach, comprised of a rumen, areticulum and an omasum, and a fourth chamber known as an abomasum.Non-limiting examples of ruminants include mammals belonging to thegenus Capra, Bos, Cervus, and Ovis.

The term “derived from,” as used herein, refers to the tissue source ororigin of an ECM. ECMs can be derived in whole or in part from tissues,such that they retain at least one component of the tissue, such as thepropria-submucosa.

The term “propria-submucosa,” as used herein, refers to a portion of theforestomach wall of a ruminant which consists of the lamina propria andthe tunica submucosa.

The term “Sealed Transmural Osmotic Flow” (STOF), as used herein, refersto a method of decellularising and/or separating the layers of tissue ororgan in which a transmural osmotic flow is established across all orpart of the wall of the tissue or organ.

The term “hypertonic,” as used herein, refers to a solution having ahigher concentration of solute relative to another solution.

The term “hypotonic,” as used herein, refers to a solution having alower concentration of solute relative to another solution.

The term “delaminated,” as used herein, refers to the separation oflayers within a tissue or organ.

The term “decellularised,” as used herein, refers to the removal ofcells and their related debris from a portion of a tissue or organ, forexample, from the ECM.

The term “breast reconstruction,” as used herein, refers to anyprocedure intended to alter the size, shape, position or appearance of abreast mound in a patient. Such procedures include, but are not limitedto, breast augmentation, mastopexy (i.e., breast lift), andreconstruction post-mastectomy.

Various aspects of the invention are described in further detail in thefollowing subsections. Unless otherwise defined, all technical andscientific terms used herein have the same meaning as is commonlyunderstood by one of ordinary skill in the art to which this inventionbelongs. In case of conflict, the present specification, includingdefinitions, will control. Although methods and materials similar orequivalent to those described herein can be used in the practice of theinvention, examples of suitable methods and materials are describedbelow. The materials, methods, and examples described herein areillustrative only and are not intended to be limiting. All publications,patent applications, patents, and other references mentioned herein areincorporated by reference in their entirety.

II. ANATOMY OF THE FORESTOMACH OF A RUMINANT

Ruminants (e.g., cattle, sheep and goats) have compound stomachs whichdiffer from the simple stomach of other mammals in that the compoundstomach is substantially larger and is divided into four sections: therumen, reticulum, omasum, and abomasum. Each of these sections has adistinct physical and histological structure. Collectively the rumen,reticulum, and omasum are known as the forestomach (proventriculus). Therumen and reticulum are intimately related in structure and function andare often referred to as rumenoreticulum. Only the last chamber of thecompound stomach, the abomasum, is structurally analogous to the simpleglandular stomach. The anatomical differences between the forestomachand the simple glandular stomach reflect their distinct functionalroles. The primary functions of the forestomach are storage,fermentation and absorption, while the simple glandular stomach performssecretory and digestive functions.

Consequently, the forestomach has gross anatomical and histologicalfeatures that are quite distinct from those of the glandular stomach.Anatomical characteristics of the forestomach are illustrated in FIG.1A, and those of the glandular stomach are illustrated in FIG. 1B.Importantly, the forestomach does not contain a glandular mucosa, but isinstead comprised of a non-glandular keratinized stratified squamousepithelium, which consists of the stratum corneum, stratum granulosum,stratum spinosum and stratum basale, and appears in many respectsanalogous to the structure of the skin. The epithelium is located on theluminal side of the forestomach, and is separated from the underlyingpropria-submucosa by a basement membrane. The abluminal side of theforestomach contains a muscle layer known as the tunica muscularis.

Stomach submucosa compositions described previously are derived from thewall of the glandular stomach, which contains the following layers: thetunica mucosa (including an epithelium layer, a lamina propria layerconsisting of reticular or fine areolar tissue, and a glandular layer),the tunica submucosa layer (composed of areolar tissue and lackingglands), the tunica muscularis layer (composed of three layers ofmuscle), and the serosa (a layer of mesothelium outside the looseconnective tissue which invests the muscle layers). The presence of theglandular layer within the stomach wall is characteristic of theglandular, gastric or simple stomach of monogastric mammals. Only thelast chamber of the complex stomach of ruminants, the abomasum, containsthis glandular layer. Additional characteristics of the forestomach andthe glandular stomach are described in Table 1.

TABLE 1 Features of the forestomach and the glandular stomach FeatureForestomach Glandular stomach Epithelium Keratinised Simple Columnarstratified squamous Glands No Yes Lamina muscularis No Yes mucosapresent lamina propria merges into Yes No submucosa creating apropria-submucosa Lamina propria Dense Loose lamina propria contains YesNo Collagen IV and Laminin Papilla on surface Yes No Lamina propriaretained Yes, dense lamina No, loose lamina propria with the propria-propria resists and glands do not resist submucosa when physicalphysical delamination epithelium is physically delamination delaminatedDense lamina propria on Yes No the luminal side of the propria-submucosaContoured surface when Yes, due to the No epithelium delaminated ECMwithin the papilla

Two unique features of the forestomach relative to the glandular stomachare that the lamina propria of the forestomach is much denser and doesnot include glands or a glandular layer. In addition, the laminamuscularis mucosa, a fine muscle layer in the basal region of the tunicamucosa layer of the glandular stomach, is absent from the rumen and mostof the reticulum. In the absence of the lamina muscularis, the laminapropria blends with the submucosa to form a layer which is collectivelyreferred to as the propria-submucosa. Also unique to the forestomach isan unusually thick and dense band of ECM within the lamina propria whichruns parallel to the epithelial surface. This band of tissue containscollagen IV and laminin, which play a critical role in cell growth,differentiation, and migration during tissue development andreconstruction. Beneath this band of tissue the ECM has a more typicalopen reticular pattern. Included in the forestomach ECM is theglycosaminoglycan heparan sulfate, an important co-factor that modulatesthe bioactivity of the growth factor FGF2. As referenced in U.S. Pat.No. 6,099,567, heparan sulfate is not present in glandular stomachsubmucosa. This is an important differentiation between the two ECMs.

Forestomach tissue also includes surface protrusions known as papillaein the rumen, reticular crests in the reticulum, and lamellae in theomasum. The propria-submucosa extends into these protrusions.

III. TISSUE SCAFFOLDS DERIVED FROM THE FORESTOMACH

According to the present invention, ECM scaffolds can be derived fromthe rumen, the reticulum or the omasum of the forestomach. Such ECMscaffolds (referred to herein as “Forestomach Matrix” or FM) arecharacterized in that they contain the lamina propria and submucosa(propria-submucosa) layers of the forestomach wall. In a particularembodiment of the invention, FM scaffolds are derived from the rumen orfrom individual laminae within the omasum. In addition topropria-submucosa, FM scaffolds may optionally include intact or partiallayers of decellularised epithelium, basement membrane, or tunicamuscularis (see FIG. 1A).

As a result of the unique structure and function of the forestomach, ECMtissue scaffolds of the invention derived from the forestomach havedifferent biochemical, structural and physical properties relative topreviously described scaffolds isolated from glandular stomach,intestine, and bladder. In particular, FM includes a dense band of ECMwithin the lamina propria. In addition, FM optionally includes an intactor fractured basement membrane. In contrast, a scaffold derived from theglandular stomach submucosa or small intestinal submucosa will includelittle if any of the lamina propria, because the lamina propria islocated mainly between the glands of the mucosa, and is consequentlyremoved as the mucosa is delaminated. Importantly, histology shows thatthe lamina propria is unusually dense, whereas the abluminal side of theFM scaffold is structured as an open reticular matrix. These differencesserve an important role in epithelial regeneration, as the dense sideacts as a barrier to cell migration, while the less dense side does notpresent a barrier and therefore allows cell invasion. This structuremakes the FM well suited for encouraging epithelial regeneration on thedense luminal side of the matrix, and fibroblast invasion on the lessdense abluminal side of the matrix, when used as a medical device fortissue regeneration. In contrast, submucosal tissue grafts derived fromthe glandular stomach and the urinary bladder have a uniform density.

The dense layer of ECM from the lamina propria contributes to theincreased thickness and strength of FM scaffolds compared to thosederived from other organs. A comparison of the thickness and burststrength of compositions derived from the forestomach and those derivedfrom other organs are provided in Examples 11 and 12.

The large surface area of the forestomach and the increased thicknessand strength of scaffolds derived from the forestomach allows theisolation of larger ECM scaffolds from the forestomach than is possiblefrom other organs. For example, ECM scaffolds of the invention can havea width as large as 5 cm (e.g., 0.5 cm, 1 cm, 2 cm, 3 cm, 4 cm, or 5cm), more preferably at least 6 cm, 7 cm, 8 cm, or 9 cm, and mostpreferably at least 10 cm or more. In addition, ECM scaffolds of theinvention can have a length as large as 5 cm (e.g., 0.5 cm, 1 cm, 2 cm,3 cm, 4 cm, or 5 cm), more preferably at least 6 cm, 7 cm, 8 cm, or 9cm, and most preferably at least 10 cm or more. Accordingly, in aparticular embodiment, FM scaffolds of the invention can have a widthand a length of 10 cm or more, a size much larger than ECM scaffoldsderived from other organs. Exemplary FM scaffolds have a surface area ofat least 100 cm², 200 cm², 300 cm², 400 cm², 500 cm², 600 cm², 700 cm²,800 cm², 900 cm², or 1000 cm² or more. In a particular embodiment, theFM scaffold has a surface area of approximately 400 cm².

Unlike scaffolds obtained from the glandular stomach, ECM scaffoldsderived from the forestomach (i.e., FM scaffolds) can include collagenIV and laminin from the basement membrane on the luminal surface.Surprisingly, these proteins are also present within the dense band ofthe lamina propria, providing important substrates for epithelial celladhesion and growth. Glandular stomach scaffolds do not typicallyinclude the epithelium or basement membrane, or portions thereof,because these layers are fragile and do not withstand physicaldelamination (see FIG. 1B). A glandular submucosal scaffold may includeremnants of the lamina muscularis mucosa on the luminal side and tunicamuscularis on the abluminal side.

FM scaffolds have a contoured luminal surface, analogous to the reteridges of the dermis. In contrast, scaffolds delaminated from smallintestine, urinary bladder and glandular stomach submucosa have arelatively smooth luminal surface. The contoured luminal surface of theFM provides a complex topology which favors epithelial regeneration.This topology is not present in ECM scaffolds derived from smallintestinal submucosa, glandular stomach submucosa or urinary bladdersubmucosa.

FM scaffolds of the invention contain important regulators of woundrepair, including but not limited to the growth factors FGF-2, TGFb1,TGFb2, and VEGF, and the glycosaminoglycans hyaluronic acid and heparansulfate. FGF2 plays an important role in wound healing by signaling cellmigration and differentiation required for the formation of new tissueand vasculature. Heparan sulphate is an important co-factor thatmodulates bioactivity of FGF2 by acting on FGF2 receptors. Heparansulphate is required for FGF2 activity and increases the stability ofFGF2. Importantly, FGF2 and heparan sulphate are not present on stomachsubmucosa. FM additionally contains fibrillar proteins includingcollagen I, collagen III and elastin, as well as adhesive proteinsincluding fibronectin, collagen IV and laminin. These proteins, inparticular collagen and elastin, contribute to the high tensile strengthand resilience of FM scaffolds. A detailed quantification of themolecular composition of FM scaffold is provided in Example 7.

In particular embodiments, FM scaffolds can be laminated together toform multi-layer sheets. For example, the laminated FM may comprise 2 ormore sheets of FM scaffold (e.g., between 2 and 30 sheets of FMscaffold, e.g., 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17,18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30 or more sheets). In aparticular embodiment, laminated FM comprises between 2 and 15 sheets ofFM scaffold, e.g., 2 to 10 sheets, 2 to 8 sheets, 2 to 6 sheets, or 2 to4 sheets of FM scaffold. The sheets of FM scaffold can be laminatedtogether using any suitable technique known in the art. In addition,lamination can be achieved using a polymer, as described below.Lamination can be achieved with or without sewing with resorbable ornon-absorbable suture material or equivalent.

The strength and physical characteristics of the FM scaffold translateinto improved handling characteristics relative to scaffolds derivedfrom thinner and weaker ECM sources. As FM scaffolds are physically morerobust than ECM scaffolds isolated from other sources, e.g., glandularstomach, they provide greater ease of handling, and are more resistantto handling deformations. This has important implications in clinicalpractice where the handling of scaffolds is necessary prior to andduring surgical procedures.

FM scaffolds can be unperforated, or they can be perforated.Perforations may be introduced into the FM scaffold using any suitablemethod, including manual drilling or laser drilling. The pore size canvary between about 10 to about 500 microns (e.g., about 10, 15, 20, 25,30, 35, 40, 45, 50, 60, 70, 80, 90, 100, 125, 150, 175, 200, 225, 250,275, 300, 325, 350, 375, 400, 425, 450, 475, or about 500 microns).Perforations may be introduced at any time during the production of FMscaffolds, but is preferably done prior to sterilization. Theperforations may pass completely through the FM scaffold, or maypenetrate only partially through the FM scaffold. In a laminated FMscaffold comprising multiple layers of FM sheets, the perforations maypass completely through all layers of the scaffold, or may pass throughonly some layers and, accordingly, penetrate only partially through thescaffold. Perforation permits cells to more easily infiltrate thescaffold, allowing for more rapid tissue ingrowth and remodeling of thescaffold.

The FM scaffolds of the invention are unique in that they combineimproved handling characteristics, are available in large formats andinclude novel combinations of biochemical constituents.

While suitable methods of isolating ECM scaffolds from the forestomachof a ruminant are described herein, the invention is intended toencompass ECM scaffolds that are isolated from the forestomach of aruminant by any means, including but not limited to the STOF method setforth below.

IV. LAMINATED TISSUE SCAFFOLDS

ECM scaffolds can be joined together to form a multi-ply laminatedsheet. Lamination of ECM scaffolds increases the strength of thescaffold, making laminated ECM compositions particularly suitable forapplications where the scaffold is required to be load bearing and/or toretain sutures or staples. Sheets may be joined together in severalorientations. For example, two or more sheets may be stacked in the sameorientation with respect to one another, i.e., such that the luminalsurface of the matrix of one sheet contacts the abluminal surface of thematrix of an adjacent sheet. In an alternative embodiment, two or moresheets may be stacked in the opposite orientation with respect to oneanother, i.e., such that the luminal surface of the matrix of one sheetcontacts the luminal surface of the matrix of an adjacent sheet, or suchthat the abluminal surface of the matrix of one sheet contacts theabluminal surface of the matrix of an adjacent sheet. Laminated FMscaffolds may be formed by bonding two or more layers of FM scaffoldtogether using a number of techniques, including, but not limited to, apolymeric adhesive layer, sewing, or simply dehydrating contacting FMlayers.

A. Laminated Tissue Scaffolds Containing Adhesive Polymers

Conventional methods of laminating ECM scaffolds to form multi-plysheets involve the use of chemical agents to crosslink the ECM scaffoldsdirectly to each other. By acting directly on the ECM scaffold, suchagents modify the scaffold and, consequently, alter the scaffold'sbiological properties. The present invention overcomes this limitationby providing methods of forming laminated ECM scaffolds (e.g., FMscaffolds) without chemical modification of the scaffold itself. Onesuch method involves distributing a polymer between one or more layersof ECM scaffold. The polymer serves as an adhesive, joining togetheralternating layers of ECM scaffold into a multi-ply composition. Thepolymer can also be applied to the outer surface(s) of an ECM scaffold.Importantly, the use of a polymer to bond a stack of ECM scaffoldsnegates the need for chemical crosslinking or other covalentmodifications of the ECM to generate a laminated composition. Thus, thebiological properties of the original ECM scaffold are retained in thelaminated scaffolds.

The methods of using polymers to generate laminated ECM compositions asdescribed herein are applicable to laminating together multiple layersof FM scaffold, or multiple layers of other ECM scaffolds known in theart, for example, ECM compositions derived from simple glandularstomach, small intestinal submucosa, bladder submucosa, or dermal ECM.In certain embodiments, a polymer can be used to form laminated sheetsof Alloderm®, Strattice®, or Surgisis®, or combinations thereof.Polymers can also be used to form a laminated composition in whichlayers of FM are laminated to other ECM scaffolds, e.g., scaffoldsderived from glandular stomach, small intestinal submucosa, bladdersubmucosa, pericardial or dermal ECM, e.g., Alloderm®, Strattice®, orSurgisis®.

In one embodiment, laminated ECM sheets (e.g., FM sheets) are formed bydistributing a polymer between two or more alternating layers of ECMscaffold (e.g., FM scaffold). The polymer may be distributedintermittently across the ECM scaffold, or it may be present as acontinuous layer. A polymer layer can be applied as intact films orsheets, or as solutions or gels. The polymer has the effect of bondingtogether two successive sheets of ECM. In a preferred embodiment, thepolymer forms a continuous and intact layer within the laminatesandwich. The polymer can additionally or alternatively be applied tothe outer surface of a laminated ECM scaffold. A range of suitablepolymers including collagen, chitosan, alginate, polyvinyl alcohol,carboxymethyl cellulose or hydroxypropyl cellulose, or combinationsthereof can be used to laminate successive sheets of ECM scaffold. Thepolymers can be applied as films, sheets, solutions, suspensions or gelsto freeze-dried sheets of ECM, then dehydrated to yield laminated ECMsheets. Alternatively, the polymers can be applied as solutions,suspensions, gels or dry films to wet ECM sheets then dehydrated toyield laminated ECM. Other suitable polymers include, but are notlimited to, the poly-glycolic acid (PGA), poly-lactic acid (PLA),Poly-lactic co-lactic acid (PLLA) and poly(lactic acid)-poly(glycolicacid) (PLGA) polymers described in any of U.S. Patent Application Nos.2002/0119180 or 2003/0031696, or U.S. Pat. Nos. 6,281,256, 6,472,210,5,885,829, 5,366,734; 5,366,733; 5,366,508; 5,360,610; 5,350,580;5,324,520; 5,324,519; 5,324,307; 5,320,624; 5,308,623; 5,288,496;5,281,419; 5,278,202; 5,278,201; 5,271,961; 5,268,178; 5,250,584;5,227,157; 5,192,741; 5,185,152; 5,171,217; 5,143,730; 5,133,755;5,108,755; 5,084,051; 5,080,665; 5,077,049; 5,051,272; 5,011,692;5,007,939; 5,004,602; 4,961,707; 4,938,763; 4,916,193; 4,898,734;4,898,186; 4,889,119; 4,844,854; 4,839,130; 4,818,542; 4,744,365;4,741,337; 4,623,588; 4,578,384; 4,568,559; 4,563,489; 4,539,981;4,530,449; 4,384,975; 4,300,565; 4,279,249; 4,243,775; 4,181,983;4,166,800; 4,137,921, the contents of which are incorporated herein byreference in their entirety.

As will be appreciated by the skilled artisan, the polymeric layercontributes to the overall performance characteristics of the laminatedscaffold. Accordingly, different strength and handling characteristicsof the laminate can be produced by altering the nature of the polymerlayer. In addition, changes in the composition of the polymer layer canbe used to alter the hydration rate of the laminate and its proteolyticstability. For example, use of a relatively hydrophobic polymer resultsin a decreased rate of hydration of the laminate, relative to a laminatecreated using a hydrophilic polymer. Non-natural and synthetic polymers(e.g., poly-vinyl alcohol) would be expected to have increased enzymaticstability relative to naturally occurring polymers, (e.g.poly-saccharide).

By distributing a polymer between successive layers of ECM scaffold,laminated compositions comprising 2 or more sheets of ECM scaffold(e.g., 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19,20, 21, 22, 23, 24, 25, 26, 27, 28, 29, or more sheets of ECM scaffold)may be generated. In an exemplary embodiment, the ECM scaffold is an FMscaffold.

In one embodiment, a gel of collagen prepared from FM may be used as thepolymer to laminate together two or more layers of FM scaffold. This gelretains the biochemical qualities of the parent FM scaffold andundergoes remodeling when used, for example, as part of a laminated FMscaffold for tissue regeneration.

B. Laminated Tissue Scaffolds Formed using Stitches or Sutures

Laminated ECM sheets (e.g., laminated FM sheets) can alternatively beformed by sewing together multiple layers of ECM. The physicalproperties of FM (size, thickness, strength, etc.) enable sheets to besewn or sutured together to form laminates for subsequent use. Laminatescan be sewn together with or without a polymer layer situated betweenindividual FM sheets.

Sewing provides another means of changing the performancecharacteristics of the laminated scaffolds. Sewing can also assistcreating devices from one or more pieces of single or laminated FMsheets that have a three-dimension architecture which is useful for aspecific anatomical site. Sewing helps to retain the three-dimensionalform of the laminate following rehydration and during handling.

Sheets of FM scaffold can be sewn using any suitable thread, includingbut not limited to absorbable suture (e.g., polyglecaprone 25(Monocryl), polydioxanone (PDS), polyglactin-910 (Vicryl), polyglycolicacid (Dexon)), non-absorbable suture (e.g., nylon (Ethilon),polypropylene (Prolene), cotton thread, or silk thread. Thread can be ofvarious thickness or gauge depending on the required strengthcharacteristics, (e.g., 6-0, 5-0 or 4-0 suture), and can be sewn using avariety of stitch lengths (e.g., 2 mm, 4 mm, or 6 mm) and patterns(e.g., straight stitch, running stitch, zig-zag stitch, overlock stitchor lock stitch).

C. Laminated Tissue Scaffolds Containing Bioactive Molecules

As described above, the invention relates in part to laminated ECMscaffolds (e.g., FM scaffolds) that include a polymer situated betweenindividual ECM sheets. This polymer helps to bind together adjacentsheets of ECM scaffold. The composition of the polymer can be modifiedin order to alter the properties of the laminated ECM scaffold,including the effect the scaffold imparts on a tissue or organ. In aparticular embodiment, the polymer can be used as a vehicle for deliveryof bioactive molecules to a tissue or organ, allowing the bioactivemolecules to be released at the site of contact. Tissue scaffoldscontaining bioactive molecules can be used, for example, to promote therate and quality of tissue regeneration, and to prevent or treat acuteor chronic infection.

Any desirable bioactive molecule can be incorporated into the polymer.Suitable molecules include, for example, small molecules, peptides orproteins, or mixtures thereof. In some embodiments, two or more (e.g.,2, 3, 4, 5, 6, 7, 8, 9, 10 or more) distinct bioactive molecules areincorporated into the polymer. Bioactive molecules can be non-covalentlyincorporated into the polymer either as suspensions, encapsulated asparticles, microparticles and/or colloids, or as a mixture thereof.Bioactive molecules can also be covalently incorporated into the polymerusing appropriate chemistries to link the bioactive molecule to thepolymer. The polymer containing one or more bioactive molecules can bedistributed between one or more ECM sheets of a laminated ECM scaffold(e.g., an FM scaffold), and/or may be applied to the outer surface ofthe scaffold. In a particular embodiment, a polymer containing a firstbioactive molecule is distributed between some layers of a laminated ECMscaffold, and a polymer containing a second bioactive molecule isdistributed between other layers of a laminated ECM scaffold. Suitablebioactive molecules include, but are not limited to, anti-microbials,analgesics, growth factors, hemostatics, pro- and anti-angiogenicagents, and combinations thereof. In particular embodiments, bioactivemolecules incorporated into a polymer layer of a laminated ECM scaffoldinclude FGF2, NGF, doxycycline, poly-L-lysine, and combinations thereof.In another particular embodiment, the polymer layer contains ananti-microbial agent and a growth factor. Importantly, as shown herein,by functionalizing the polymer layer of an ECM scaffold laminate with abioactive molecule, the inherent biological properties of the ECMscaffold are not altered, in contrast with conventional methods in whichadditional molecules are incorporated into ECM scaffolds by attachmentto the ECM directly.

Bioactive laminated ECM scaffolds (e.g., FM scaffolds) in which thebioactive agent is incorporated in an adhesive polymer (e.g., in apolymer layer) situated between successive layers of ECM have severalcritical advantages over alternative compositions in which a bioactiveagent is conjugated directly to the ECM. Firstly, incorporation of thebioactive agent into a polymer or polymer layer does not change theinherent composition of the ECM scaffold, as it does not requirecovalent or chemical modification of the scaffold to achieve laminationor loading with the bioactive agent. Secondly, the use of apreformulated bioactive polymer layer as a vehicle for delivery of abioactive agent allows greater control and consistency in the uniformityof dosing of the bioactive molecule.

IV. USES OF FORESTOMACH MATRIX COMPOSITIONS

Forestomach matrix scaffolds are well suited to a wide range of tissueregeneration applications. They can be used to cover tissue deficitssuch as wounds, and to reinforce and/or repair soft tissue. They can beused as single or laminated sheets, and may be formed into customizedconforming devices to suit particular organs, anatomical sites, orspecific surgical applications. FM scaffolds are secured in place usingany suitable method known in the art, including sutures, staples ordressings.

In one embodiment, the FM scaffolds are used to cover extensivetraumatic wounds or burn injuries, overcoming the inconvenience andcomplexity of joining a number of smaller scaffold devices together toachieve full coverage. Accordingly, particular FM scaffolds of theinvention can cover a wound or injury having a width of 10 cm or moreand a length of 10 cm or more.

The presence of the dense layer of connective tissue within the laminapropria and the contoured surface topology make FM well suited in dermaland epithelial regeneration applications. The high tensile strength ofFM scaffolds is also particularly useful where the scaffold is requiredto load bear or is placed under tension. Accordingly, in a particularembodiment, the FM scaffolds of the invention are applied to wounds andsurgical sites where there is a need to stimulate tissue repair orregeneration or provide tissue reinforcement. The ECM is naturallyremodeled over time, such that FM scaffolds are resorbed and replaced byhost tissue.

In other embodiments, FM scaffolds of the invention are used to replacedamaged, diseased, or missing heart valves, arteries, veins, urinarybladder, liver, portions of the gastrointestinal tract, or as templatesfor repair or replacement of head and neck structures. FM, in any of anumber of its solid or fluidized forms, can be used as a scaffold fordermal or epidermal repair, injected into various body sphincters suchas urinary sphincter or esophageal or gastric sphincters, folded into atube or partial tube as a conduit for the restoration of nervous tissueor extruded or molded into any shape suitable for its application as atissue regenerative composition. Accordingly, the FM scaffolds of theinvention can be sutured into place in solid sheet form, placed inwounds or body locations in a gel form, or injected in its liquid orparticulate form. FM scaffolds of the present invention induce growth ofendogenous tissues including epithelial and connective tissues whenplaced in contact with target tissues in vivo. In addition, the FM canoptionally be combined with cells to create tissue constructs togenerate new skin, cardiovascular, urogenital, neurological, fascia,tendons, sheaths, ligaments and gastrointestinal tissues. FM can also beseeded with keratinocytes on the denser luminal side of the matrix, andwith fibroblasts on the less dense abluminal side for use in certaindermatological applications. FM scaffolds can be seeded with a varietyof cell types, including stem cells, for applications in regenerativemedicine.

In still other embodiments, FM scaffolds of the invention serve as asubstrate for attachment in in vitro cell culture and as a scaffold forcell growth in tissue engineering applications, where FM can promoteproliferation and/or induce differentiation of eukaryotic cells.Protocols utilizing non-FM submucosal tissue in in vitro cell cultureapplications are described, for example, in U.S. Pat. No. 5,695,998,incorporated herein by reference in its entirety. These methods aregenerally applicable to the use of FM as a substrate for promoting invitro cell culture. In general, this involves contacting FM witheukaryotic cells in vitro under conditions conducive to eukaryotic cellgrowth. As described herein, FM scaffolds of the invention can also beused for constructing devices for drug delivery.

As described herein, FM scaffolds can increase proliferation of cellslocalized near the scaffold attachment site. Accordingly, FM scaffoldscan be used to promote, stimulate, or increase cell proliferation in atissue or organ. In a preferred embodiment, FM scaffolds are used topromote, stimulate, or increase cell proliferation within a woundedtissue or tissue deficit, e.g., within a regenerating wound.

FM scaffolds also promote vascularization (e.g., angiogenesis) within atissue or organ to which the FM scaffold adheres. Accordingly, FMscaffolds can be used to promote, stimulate, or increase vascularizationof a tissue or organ. In a preferred embodiment, FM scaffolds are usedto promote, stimulate, or increase vascularization of a wounded tissueor tissue deficit, e.g., within a regenerating wound. Improvingvascularization is one way in which FM scaffolds promote wound closureand improve the quality of wound healing.

Bioactive laminated FM scaffolds described herein have many additionalclinical applications, including but not limited to delivery ofantibiotics (e.g., amoxicillin, penicillins, poly-amines, or quinolines)to surgical sites to treat, inhibit or prevent microbial infection;delivery of antibiotics to wounds and tissue deficits to treat, inhibitor prevent microbial infection at the site; delivery of growth factors(e.g., FGF2, VEGF or PDGF) to a wound or surgical site to promote tissueregeneration and/or vascularization of the tissue; delivery of enzymaticinhibitors to reduce proteolytic activity in chronic wounds; delivery ofnitric oxide analogs to a wound or surgical site to promote tissueregeneration; delivery of antimicrobials or anti-biofilm agents to awound or surgical site to inhibit or prevent infection and/or formationof biofilms.

FM scaffolds of the invention may be formulated and used in a variety offormats, including but not limited to powder, emulsion (fluidized FM),gel or extract. Moreover, the FM scaffolds may be sterilized prior touse by conventional methods, including ethylene oxide treatment, gammairradiation treatment, gas plasma sterilization, or e-beam treatment.

V. LAMINATED FM SCAFFOLDS USEFUL FOR BREAST RECONSTRUCTION

The characteristics of FM tissue scaffolds of the invention, e.g.,strength, elasticity, suture retention, etc., as described herein, makethem suitable for a variety of applications in which there is a need tosupport or reinforce soft tissue. In a particular embodiment, the FMtissue scaffolds are used to cover, position and/or secure breastprosthetics during breast reconstruction, or to cover, position and/orsecure native breast tissue or breast prosthetics during mastopexy(i.e., “breast lift”).

Breast augmentation is a popular cosmetic procedure in which aprosthesis, i.e., a breast implant, is typically positioned in the chestin one of three positions: over the pectoralis major muscle and underthe breast tissue (subglandular), partially under the muscle (partialsubmuscular), or completely under the muscle (submuscular). Regardlessof the location of the implant, the aesthetic outcome of the proceduredepends largely on the ability of the surrounding tissue to support theprosthesis such that the prosthesis maintains its position within thepatient. Over time, prosthetics can be displaced medially, resulting insymmastia; laterally, resulting in implant excursion into the axilla ofthe chest cavity, or inferiorly, resulting in “bottoming out.” Afrequent cause of malposition is inadequate soft tissue support for theweight of the implant. Inadequate soft tissue support is common inpatients who receive very large implants, for example, and in patientswho have lost a large amount of weight.

This problem is exacerbated in patients undergoing breast reconstructionfollowing treatment for breast cancer, in particular. Cancer treatmentssuch as radiation or chemotherapy weaken the soft tissue needed tosupport the prosthesis. In addition, attaining sufficient muscle or softtissue coverage of the prosthesis following mastectomy is a difficulttechnical challenge. The feasibility of attaining adequate coverage isdependent on the extent of tissue loss and the quality of the remainingtissue. When adequate coverage is not possible, coverage isconventionally achieved by transferring muscle tissue from another siteon a patient, which can be associated with donor site morbidity, poorhealing, scarring and contracture, and the risk of infection andpotential flap necrosis.

The FM tissue scaffold of the present invention addresses these problemsin that it can reinforce breast tissue and achieve adequate support fora variety of breast prostheses during breastaugmentation/reconstruction. The tissue scaffold may also be used tosupport native breast tissue or breast prosthetics during mastopexy. Theadded support of the tissue scaffold is provided, in part, byextracellular matrix derived from the forestomach of a ruminant, asdescribed herein.

FM scaffolds for breast reconstruction can, for example, be flat, orhave a concave shape. In a preferred embodiment, the FM scaffold hasconcavity. Such concavity provides improved conformity and approximationto the rounded shape of the breast tissue and/or the breast prosthesis,reducing dead space and improving positioning, tissue apposition andfixation as compared to a flat sheet. FM is well suited to thisapplication because the natural curvature and shape of the forestomachare useful for forming ECM scaffolds having a natural concavity. Inaddition, FM scaffolds can be formed around a mold to alter the shape asneeded for particular applications (e.g., by increasing or decreasingthe curvature of the scaffold).

FM scaffolds for breast reconstruction also can comprise a single orlaminated sheet of FM. For example, multiple sheets of FM can belaminated together to increase the strength and thickness of thescaffold, as described herein. In order to support a breast prosthesisand/or native breast tissue, a relatively strong scaffold is needed.Accordingly, FM scaffolds for breast reconstruction can advantageouslyinclude a laminated sheet containing 2 or more sheets of FM joinedtogether (e.g., 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15 or moresheets). The sheets of FM may be laminated using polymers, usingstitches or sutures, or using both polymers and stitches or sutures, asdescribed herein. In a particular embodiment, the layers of laminated FMscaffolds for breast reconstruction are secured together by stitching.Use of stitching and/or adhesive polymers for lamination of multiple FMsheets ensures that the FM scaffold retains its three-dimensional shapeprior to and during implantation in a patient.

The FM scaffold also may be perforated or unperforated, as describedherein. Perforation reduces the risk of fluid accumulation and seromaformation under the implant, and permits cells to more easily infiltratethe scaffold, allowing for more rapid tissue ingrowth and remodeling ofthe scaffold.

An FM scaffold for breast reconstruction may have a number of differentshapes in order to adequately provide coverage for a breast prosthesisduring breast augmentation, or for native breast tissue duringmastopexy. In preferred embodiments, the FM scaffold has a crescentshape, as illustrated in FIG. 2(A), or an elliptical shape, asillustrated in FIG. 2(B). A semi-circle or half-moon shape may also beused. In one embodiment, the scaffold is sufficiently large tocompletely or partially cover the lower and/or lateral sections of thebreast prosthesis or breast tissue. This allows the scaffold to supportthe lower pole of the breast prosthesis and/or native breast tissue,emulating the inferior and lateral mammary folds. In this embodiment,the scaffold may be placed in a horizontal or vertical orientation. Thescaffold can also be sized for placement in a vertical orientation onthe lateral or medial side of a breast prosthesis, to inhibit lateral ormedial displacement of the prosthesis.

In particular embodiments of the invention, the FM scaffold for breastreconstruction is between about 3 cm to about 35 cm in length (e.g., 3,4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22,23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, or 35 cm), and about 3cm to about 35 cm in width (e.g., 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13,14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31,32, 33, 34, or 35 cm). An exemplary FM scaffold for breastreconstruction is about 3 cm to about 12 cm wide and about 25 cm toabout 35 cm long. Additional suitable shapes and sizes of scaffolds areknown in the art, as described, for example, in WO 2008/016919 and US2007/0088434 A1, the contents of which are incorporated herein byreference in their entirety. These shapes and sizes all can be appliedto FM scaffolds of the present invention.

When implanted into a patient, the FM scaffold for breast reconstructionundergoes controlled biodegradation as a result of living cellreplacement, such that the original scaffold is remodeled. Infiltrationof a patient's cells into the scaffold ultimately results in replacementof the ECM of the scaffold with matrix produced by the infiltratingcells. Over time, the scaffold undergoes normal constructive remodeling,resulting in the formation of new tissue to support the breastprosthesis and/or breast tissue. This approach eliminates the need fortransferring tissue from another site on the patient, especially in thecase of post-mastectomy reconstruction, thereby reducing the complexityand duration of the surgical procedure.

Accordingly, FM scaffolds for breast reconstruction described herein maybe used in a wide range of procedures involving breast augmentation ormastopexy, including, for example, in a prophylactic nature at the timeor initial placement of a breast prosthesis, in subsequent correctiverepositioning procedures, in post-mastectomy reconstruction, and inbreast lift procedures. Methods for using tissue scaffolds in theforegoing procedures are well known in the art. Such methods typicallyinvolve fixation of the scaffold in the desired position, e.g., acrossthe lower and lateral sections of the breast to support the lower poleof a breast prosthesis/breast tissue, or on the lateral or medial sideof the breast to inhibit lateral or medial displacement. Fixation may beachieved using any suitable method known in the art, for example, byplacement of sutures or staples, or with use of a tacking device. Overtime, the FM scaffold becomes integrated with the surrounding tissue.Other exemplary techniques for using artificial tissue scaffolds inbreast reconstruction are described, for example, in WO 2008/016919 andUS 2007/0088434 A1, the contents of which are incorporated herein byreference in their entirety. Such techniques are likewise applicable tothe FM tissue scaffolds of the invention in breast reconstructionprocedures.

VI. ISOLATING TISSUE SCAFFOLDS FROM MAMMALIAN ORGANS USING SEALEDTRANSMURAL OSMOTIC FLOW (STOF)

Extracellular matrix scaffolds are traditionally produced fromgastrointestinal tissue (e.g., glandular stomach & small intestine) orurogenital tissue (e.g., bladder) by removing the epithelial cells andmuscularis mucosa either physically or by chemical separation before orafter the tissue is soaked in processing solutions. Physical separationof the mucosal layer typically removes the epithelium, basementmembrane, glandular layer, and most if not all of the lamina propna andtunica muscularis.

The decellularisation process removes antigenic components from thescaffold, while conserving the biological activity as well as themechanical and structural integrity of the ECM. Physical methods of cellremoval conventionally include snap freezing, mechanical force,agitation, and sonication. Chemical agents conventionally used fordecellularisation include alkaline and acid treatments, non-ionicdetergents (e.g., Triton X-100), ionic detergents (e.g., SDS, TritonX-200), zwitterionic detergents (e.g., CHAPS, SB-10, SB-16),Tri(n-butyl) phosphate, hypotonic and hypertonic treatments, andchelating agents (e.g., EDTA, EGTA). Enzymatic processing methodsconventionally use trypsin, dipase, endonuclease, and exonuclease.

Processing methods used to date immerse either the open intact organ ora section of the organ in a sequence of processing solutions, therebyexposing all surfaces to each of the solutions. The differentcharacteristics of muscle and epithelial tissue can mean that a solutionwhich has a positive effect on one tissue layer may have a negativeimpact on another tissue layer. This often results in the use ofmultiple processing steps, extending the processing time. All processingsolutions will have an impact on the biochemical composition, tissueultrastructure, and mechanical behavior of the remaining ECM. Shorterprocessing times are desirable because prolonged exposure to processingagents, for example, Triton X-100, SB-10, SB-16, Triton X-200, SDS, andtrypsin can be detrimental to the ECM. Shorter processing times alsoallow higher throughput, improving processing economics and reducingexposure to cellular proteases, which can damage the native ECMultrastructure. Processing methods used to date rely on diffusion of theprocessing solutions through the tissue, which can be enhanced somewhatwith agitation and increased temperature. Accordingly, processing tissueat physiological temperatures sholtens processing time with respect toprocessing at 4° C. However, physiological temperatures increaseendogenous protease activity and, depending on the solution used, canencourage the growth of microbial contaminants.

The present invention overcomes these obstacles and provides an improvedmethod for producing ECM scaffolds from mammalian tissues or organs.According to the method of the invention, different solutions areisolated on each side of a tissue, allowing each solution to beoptimized to target the respective tissue layers. One solution isprepared such that it is hypertonic or hypotonic with respect to anothersolution, such that when the solutions are exposed to opposite sides ofa tissue or organ (e.g., the luminal and abluminal sides), a transmuralosmotic flow through the wall of the tissue or organ is established.This method of processing aids in decellularisation, separation oftissue layers, and removal of cell debris and processing agents. Inaddition, this method effectively reduces processing times.

Methods of the invention described herein, in which a transmural osmoticflow is established across the wall of an organ, are referred to as“Sealed Transmural Osmotic Flow” (STOF). STOF methods can be used toprocess any intact or sealed animal or human tissues to separate and/ordecellularise tissue layers, and to thereby extract ECM based scaffolds.Accordingly, this method may be used to process the forestomach tissueof a ruminant to produce FM scaffolds, or to process any other mammaliantissue. An exemplary, non-limiting embodiment of the STOF method isillustrated in FIG. 3. This figure depicts an organ that has been filledwith one solution, sealed and then immersed in another solution. Thedifference in salinity between the two solutions results in a transmuralosmotic flow. It will be appreciated that an osmotic gradient can beestablished in either direction by changing the placement of thesolutions (i.e., hypertonic and hypotonic solutions). The gradient ispreferably established in a direction mimicking the natural flow of theorgan. For example, when processing tissue from the forestomach of aruminant, the gradient is preferably established from the luminal to theabluminal surface of the tissue. Using biomimicry to determine thedirection over which to establish the transmural flow allows the naturalphysiology and flow properties of the tissue to enhance processing.

The STOF method of the invention is particularly useful for processingtissues that are difficult to delaminate, such as those having akeratinized stratified squamous epithelium, because it allows tissuelayers to be targeted with specific agents. STOF can be used to processany intact or partially intact tissue or organ which can be scaled suchthat a transmural osmotic flow may be created. This includes, forexample, tissues or organs of gastrointestinal, urogenital,cardiovascular and dermal origin. This approach may be used to process awide range of tissues or organs using various processing solutions,including but not limited to those containing salts (e.g., NaCl, KCl,EDTA, EGTA), detergents (e.g., Triton X-100, Triton X-200, sodiumdodecyl sulfate, sodium deoxycholate, CHAPs, sulfobetaine, tri(n-butyl)phosphate) and enzymes (e.g., trypsin, endonucleases and exonucleases).Accordingly, this method can be adapted for use with any animal or humantissue or organ which can be fashioned so that the tissue or organ, or aportion thereof, is sealed to allow the creation of a transmural osmoticgradient. A transmural osmotic gradient may also be established across asection of tissue which isolates one solution in a container from asecond solution in which it is immersed.

The STOF method performs very well at low temperatures, because the flowis driven by an osmotic gradient rather than being reliant on diffusionand Brownian motion. The rapid efficiency of this method at lowtemperatures is particularly advantageous in that it minimizes intrinsicprotease degradation of biological components of the ECM scaffold, andalso prevents microbial growth. In one embodiment, the method isperformed at a temperature of less than 6° C. (e.g., 2° C.-4° C.) inless than 36 hours (e.g., preferably in 24 hours or less). This methodhas been used effectively, for example, to delaminate the keratinizedstratified squamous epithelium of the forestomach of a sheep at 4° C. inless than 24 hours. By comparison, processes for generating an esophagusacellular matrix scaffold have been previously reported which takelonger than a week (see, for example, Bhrany et al., “The Development ofan Esophagus Acellular Matrix Tissue Scaffold,” Tissue engineering(2006) 12(2), incorporated herein by reference). This improvement ispossible because the transmural osmotic flow draws the processingsolutions through the tissue. In another embodiment, the method isperformed at or near room temperature (e.g., 18° C.-24° C.). At or nearroom temperature, processing times can be further reduced. For example,the STOF process has been used to decellularize and delaminate ovineforestomach at room temperature in less than 6 hours. The STOF processalso imparts the benefit of treating the different tissue surfaces withdifferent solutions that are optimized to remove the targeted layer(e.g., muscle or epithelium).

The transmural osmotic flow properties are such that the amount of fluidthat passes through a tissue is dependent on surface area. The surfacearea of the tissue may be controlled by the amount of liquidencapsulated within it (the level of distension). Accordingly, a tissueor organ may be fully or partially distended to further increase thesurface area of the tissue exposed to the osmotic gradient, and therebyincrease the osmotic flow passing into the tissue or organ.

The STOF method of the invention may be used to remove all or a part ofa single layer or multiple layers of the tissue being processed. Forexample, all or a part of the epithelium, basement membrane or tunicamuscularis, and combinations thereof, may be removed during or followingprocessing. A number of different solutions may be employed, dependingon the tissue to be processed and the composition of the tissue layersthat are to be delaminated. In one embodiment, a hypertonic solution isencapsulated within a tissue or organ, and the tissue or organ isimmersed in a hypotonic solution. In another embodiment, a hypotonicsolution is encapsulated within the organ, and the organ is immersed ina hypertonic solution. In both cases, the direction of osmotic flow willbe from the surface of the organ in contact with the hypotonic solutionto the surface in contact with the hypertonic solution.

The hypertonic solution may contain one or more buffers, detergents,salts or combinations thereof. Likewise, the hypotonic solution maycontain one or more buffers, detergents, salts, or combinations thereof.In all instances, the hypertonic solution contains a higherconcentration of solute than the hypotonic solution. For example, ahypertonic salt solution contains a higher concentration of salt than ahypotonic solution. In a preferred embodiment, the hypertonic solutioncontains a higher concentration of salt than the organ being processed,and the hypotonic solution contains a lower concentration of salt thanthe organ being processed.

In a particular embodiment, the hypertonic solution contains NaCl.Suitable NaCl concentrations range, for example, between about 0.5 M-10M (e.g., about 0.5 M, 1 M, 1.5M, 2M, 2.5M, 3M, 3.5M, 4M, 4.5M, 5M, 5.5M,6M, 6.5M, 7M, 7.5 M, 8 M, 8.5 M, 9 M, 9.5 M or 10 M. In an exemplaryembodiment, the hypertonic solution contains about 4 M NaCl.

In another particular embodiment, the hypotonic solution contains TritonX-200 and EDTA. Suitable concentrations of Triton X-200 in the hypotonicsolution range, for example, between about 0.001% and 1% (e.g., about0.001%, 0.005%, 0.01%, 0.05%, 0.1%, 0.25%, 0.5%, 0.75%, or 1%). Suitableconcentrations of EDTA in the hypotonic solution range, for example,between about 0.01% and 1% (e.g., about 0.01%, 0.05%, 0.1%, 0.25%, 0.5%,0.75%, or 1%). In an exemplary embodiment, the hypotonic solutioncontains about 0.028% Triton X-200 and about 0.1% EDTA.

In another particular embodiment, the hypotonic solution contains SDS.Suitable concentrations of SDS in the hypotonic solution range betweenabout 0.001% and 1% (e.g., about 0.001%, 0.005%, 0.01%, 0.05%, 0.1%,0.25%, 0.5%, 0.75%, or 1%). In an exemplary embodiment, the hypotonicsolution contains about 0.1% SDS.

In another exemplary embodiment, the hypotonic solution contains about0.028% SDS. In a particular embodiment, the hypotonic solution containsabout 0.028% Triton X-200, 0.1% EDTA, and 0.1% SDS. In anotherparticular embodiment, the hypotonic solution contains about 0.028%Triton X-200, 0.1% EDTA, and 0.028% SDS.

After initial processing of an organ is achieved by encapsulation of onesolution within the organ, followed by immersion of the organ in asecond solution, such that a transmural osmotic flow through the wall ofthe organ is established, further processing is possible (but notrequired) by removal of the organ from the second solution, andimmersion of the organ in a third solution, such that a transmuralosmotic flow is again established. The direction of transmural osmoticflow is preferably in the same direction during both stages ofprocessing (i.e., from the luminal to the abluminal surface, or from theabluminal to the luminal surface). The solution encapsulated within theorgan need not be removed prior to or during further processing.

Accordingly, in an exemplary embodiment, a hypertonic solutioncontaining NaCl (e.g., about 4 M NaCl) is encapsulated within amammalian organ (or a portion thereof). The organ is then immersed in ahypotonic solution containing Triton X-200 and EDTA (e.g., about 0.028%Triton X-200 and 0.1% EDTA). The organ is then immersed in a hypotonicsolution containing SDS (e.g., about 0.1% SDS or about 0.028% SDS). Thesolutions are selected based on the desired effect they impart on theprocessed tissue. For example, a hypotonic solution containing TritonX-200 and EDTA targets the breakdown of the basement membrane betweenthe epithelium and propria submucosa, while a hypotonic solutioncontaining SDS is primarily used to achieve decellularisation. Use ofthe SDS solution in a STOF method speeds up the removal of cell wallsand cellular debris. Accordingly, multiple rounds of STOF processing maybe used to achieve the benefit imparted by multiple distinct processingsolutions.

In another exemplary embodiment, the STOF method is used to separate anddecellularise the tissue layers of the forestomach of a ruminant togenerate FM. The process may be used, for example, on the forestomach ofcattle, goats or sheep. FM can be generated from the rumen, thereticulum or the omasum using the STOF method. In an exemplaryembodiment, the method is used to generate FM from the forestomach ofsheep (Ovis aries). The forestomach's epithelial surface is akeratinized stratified squamous epithelium which is tightly bound to abasement membrane. It is adapted to resist damage from abrasion, makingit difficult to remove using other manual or mechanical methods. In apreferred embodiment, a transmural osmotic flow is established acrossthe forestomach from the luminal to the abluminal surface, mimicking thenatural flow of the organ.

In another exemplary embodiment of the invention, three particularsolutions are employed for use in forestomach tissue separation and cellremoval in a timely manner using the STOF method. Solution A containsabout 4 M NaCl, Solution B contains about 0.028% Triton X-200 and 0.1%EDTA, and Solution C contains about 0.1% SDS. Solution A is hypertonicwith respect to Solution B and Solution C. Using these solutions toprocess forestomach tissue, the inverted forestomach is filled withapproximately 10 L of Solution A and can be sealed with a cable tie. Thefilled rumen can then be immersed in Solution B for approximately 16hours. This combination targets the breakdown of basement membranebetween the epithelium and the propria submucosa. The transmural osmoticflow and exposure of the muscle layer to Solution A additionally causesthe muscle fibers to soften, aiding physical separation. The filledforestomach can then be immersed in Solution C for approximately 4 hoursto achieve decellularisation.

The STOF method speeds up the removal of cell walls and cellular debris.Using the protocol condenses the processing timeframe with respect totraditional methods of processing, as flow is not simply limited toBrownian motion. Moreover, the process can be performed at lowtemperature (e.g., 4° C. or less) to limit the activity of endogenousproteases and microbiological growth. The ability to process the tissueat low temperatures and in short timeframes enables recovery of higherlevels of biological molecules native to the ECM (e.g., growth factors,fibrillar proteins, adhesive proteins, glycosaminoglycans, etc.) in thefinished scaffold with respect to processing using other methods.Accordingly, an ECM scaffold generated using the STOF method hasdistinct biochemical characteristics with respect to scaffolds isolatedby other processes.

The present invention is further illustrated by the following examples,which should not be construed as limiting.

EXAMPLES Example 1 Preparation of Forestomach Matrix (Fm) by SealedTransmural Osmotic Flow (STOF)

Forestomachs from lambs less than two years old were sourced from alocal abattoir. The forestomach was inverted so that the epithelialsurface was on the outside and the muscular layer was on the inside. Itwas filled so that the organ was distended with 10 L of 4 M NaCl andsealed with a cable tie. The forestomach was then suspended in asolution containing 0.1% EDTA and 0.028% Triton X-200 for 16 hours,after which it was transferred into a solution containing 0.1% SDS for afurther 4 hours. Care was taken to ensure that the organ does notcontact other surfaces, including one another, which can limit osmoticflow. This process allowed the Triton X-200/EDTA and SDS solutions to bedrawn through the tissue, causing decellularisation and effectivelydisrupting the basement membrane.

The forestomach was removed from the STOF process and the contents wereemptied. It was then cut so as to open the organ such that the muscularand epithelial surfaces could be delaminated. This may be carried outby, for example, either manual or mechanical scraping in order toisolate the FM. The FM was washed for 30 minutes using water at roomtemperature with stirring and then was transferred to 0.1% peraceticacid, 1 M NaCl, in 5% ethanol. This was mixed for 60 minutes followed by4 sterile wash steps in water (15 min), PBS pH 7.2 (15 min), water (15min), and PBS pH 7.2 (15 min). The FM was then freeze dried, packagedaseptically and terminally sterilized with ethylene oxide. Thisembodiment of the STOF process is outlined in FIG. 4. Histologicalexamination of Haematoxylin and Eosin (H&E)-stained sections confirmedthat decellularisation was effective. This was further verified bycomparing the concentration of DNA in forestomach tissue and FMscaffold. Papain digested samples were incubated with Hoechst 33258 dye(10 μg/mL, Sigma—Missouri, USA), and relative fluorescence units werequantified using a microtitre plate reader. Total DNA concentration wascalculated from a standard curve of calf thymus DNA (Sigma—Missouri,USA). As shown in FIG. 5, nucleic acid content was significantlydecreased in the FM scaffold as compared to the forestomach tissue,indicating that the FM scaffold was decellularised.

Example 2 Flow Properties and the Effect of Tissue Orientation duringthe STOF Method

The following experiment was performed to determine the relationshipbetween the surface area of the forestomach and the osmotic flow ofwater passing into the forestomach. In this study, 4 M NaCl and waterwere used to establish an osmotic gradient across forestomach tissue.The surface area of the forestomach can be approximated based on thesurface area of a sphere (4 pr²). The volume of the liquid in theforestomach before and after STOF was used to calculate the volumechange of the interior solution (L). There was a linear correlationbetween the surface area of the forestomach (degree of distension)before STOF and the flow of water into the forestomach (volume change),as depicted in FIG. 6. As shown therein, the osmotic flow of waterpassing into the forestomach increased with increasing surface area ofthe sealed forestomach. A reduction in osmotic flow rate was typicallyobserved over time, with the greatest flow achieved during the first 24hours of STOF.

As a means of evidencing the flow through the forestomach along anosmotic gradient during STOF processing, the weight of forestomachs wasmeasured before and after the transmural osmotic flow was applied. TheSTOF setup was similar to that described in Example 1 and FIG. 3. Theweight of the forestomach increased during STOF as a consequence offluid moving into the tissue. Forestomachs at the beginning of theprocess weighed 789+/−45 g, whereas at the end of the process the sameforestomachs weighed over 1160+/−92 g. Fluid passing into theforestomach tissue resulted in weight gain as tissue hydrationincreased. These results are depicted in FIG. 7.

Tissue orientation significantly impacted the results achievable withSTOF. Establishment of an osmotic gradient across the forestomach in thedirection of natural, physiological flow (i.e., flow from epithelium tomuscle, e.g., from the luminal surface to the abluminal surface of theforestomach) resulted in successful removal of the muscle layer,epithelial layer and cells from the forestomach. In contrast, when anosmotic gradient was established against direction of naturalphysiological flow (i.e., flow from muscle to epithelium, e.g., from theabluminal surface to the luminal surface of the forestomach) the tissuebecame dehydrated and tacky, and subsequent removal of muscle andepithelial layers was more difficult. This data is summarized in Table2. The direction of flow also affected the appearance of the tissue atthe end of processing.

TABLE 2 Tissue orientation and flow properties Tissue Interior InteriorExterior Exterior appearance Ease of tissue layer solution tissue layersolution Transmural flow after STOF delamination Muscle 0.1% Epithelium4 M Muscle→Epithelium Dehydrated Poor SDS NaCl Muscle 4 M Epithelium0.1% Epithelium→Muscle Hydrated Good NaCl SDS Epithelium 0.1% Muscle 4 MEpithelium→Muscle Hydrated Good SDS NaCl Epithelium 4 M Muscle 0.1%Muscle→Epithelium Dehydrated Poor NaCl SDS

Example 3 Preparation of Laminated Forestomach Matrix Using AdhesivePolymers

The FM scaffolds described herein may be formatted as single sheets, ormay be laminated to form multi-ply laminated compositions of FM. Thefollowing example describes the preparation of a laminated FM scaffoldusing either a collagen polymer adhesive or an adhesive layer preparedfrom polyvinyl alcohol or hydropropyl cellulose. To prepare the collagenbonded laminate, a gel of collagen polymer was firstly prepared from theparent FM scaffold. This collagen gel retained the same biochemicalqualities as the parent material and undergoes remodeling when used aspart of a laminated FM scaffold in tissue regeneration. The collagenpolymer gel was prepared using heat denaturation of the parent FMscaffold. Finely powdered FM (10% w/v) was heated at 95° C. in purifiedwater or PBS for 90 minutes. The mixture was centrifuged at 10 k rpm for30 minutes at 35° C. to remove particulates, and the supernatant wasretained. On standing and cooling to room temperature the solutionsolidified to a gel. The gelation of the collagen suspension wasreversible, and as such, was heated (>37° C.) prior to directapplication as a gel to FM sheets as a layer between two (or more)sheets. On cooling and drying the collagen gel effectively laminated theFM sheets to produce 2-ply laminates.

As the polymeric layer contributes to the overall performancecharacteristics of the laminated scaffold, different strength andhandling characteristics of the laminate were produced by altering thenature of the polymeric adhesive layer. Additionally, changes in thepolymeric layer were used to alter the hydration rate of the laminate,its proteolytic stability, and for the delivery of bioactives.

Polyvinyl alcohol and hydroxypropyl cellulose polymeric adhesive wereapplied as dry films which rehydrated on contact with the wet FMscaffold. Subsequent freeze-drying of the polyvinyl alcohol andhydroxypropyl cellulose containing laminates dehydrated the polymericlayer to yield a laminated sandwich.

The 2-ply laminates were tested for uniaxial strength according toExample 13. Results are summarized as Table 3. Polyvinyl alcohol andhydroxypropyl cellulose laminated FM scaffolds were significantlystronger than collagen laminated FM scaffolds. However, increasedstrength was offset by a decrease in the elasticity of polyvinyl alcoholand hydroxypropyl cellulose laminated FM scaffolds, as reflected by anincrease in the modulus of elasticity and a decrease in the maximumelongation. Polyvinyl alcohol laminated FM scaffold performed best basedon yield stress, a term normalized to the thickness of the sample.

This example demonstrates that as the polymeric layer contributes to theoverall performance characteristics of the laminated scaffold, differentstrength and handling characteristics of the laminate were produced byaltering the nature of the polymeric adhesive layer.

TABLE 3 Biophysical characterization of FM scaffold laminated withcollagen, polyvinyl alcohol and hydroxypropyl cellulose polymericadhesive layers. Maximum Modulus of Maximum Tangential Elasticity YieldThickness Maximum Elongation Stiffness (Youngs′) Stress (mm) Load (N)(mm) (N/mm) (GPa) (MPa) Collagen 0.306 ± 0.018 17.945 ± 3.088 19.418 ±0.721 1.347 ± 0.262 0.05 ± 0.01  9.77 ± 1.68 Hydroxypropyl 0.363 ± 0.02323.533 ± 4.000 12.536 ± 0.382 2.790 ± 0.438 0.096 ± 0.016 10.80 ± 1.84cellulose Polyvinyl 0.247 ± 0.012 22.292 ± 1.172 13.502 ± 0.222 2.385 ±0.152 0.121 ± 0.01  15.07 ± 0.79 alcohol Errors represent standarderrors from five experiments.

Example 4 FM Scaffolds Sewn Into Laminates

The FM scaffolds described herein have intrinsic physical properties(e.g. size, thickness, strength) that enable FM scaffold sheets to besewn or sutured together to form laminates. Laminates can be sewn orsutured together with or without a polymer situated between individualFM sheets. The following example describes the preparation of a 2-plylaminated FM scaffold by stitching the FM layers together with cottonthread.

FM scaffolds were sewn together using cotton thread to form 2-plylaminates. A sewing machine was used to create a straight stitch (approx2 mm stitch length) through two layers of FM scaffold sheets. The 2-plylaminate was stitched together using parallel stitches approximately 5mm apart and running the length of the laminate. Uniaxial strengthtesting of the sewn laminates was conducted using the protocol describedin Example 13, and the sewn laminates were compared with 2-ply laminatescreated using a layer of polymeric collagen adhesive that had not beensewn. Results are shown in Table 4. The strength of the sewn laminateoutperformed the collagen-based laminate, as described by the maximumload and yield stress. However, as strength increased, elasticity andelongation of the sewn laminate was reduced relative to thecollagen-based laminate.

TABLE 4 Biophysical characterization of FM scaffold laminated withcollagen, compared with a sewn FM scaffold laminate. Maximum Modulus ofMaximum Tangential Elasticity Yield Thickness Maximum ElongationStiffness (Youngs′) Stress (mm) Load (N) (mm) (N/mm) (GPa) (MPa)Collagen 0.306 ± 0.018 17.945 ± 3.088 19.418 ± 0.721 1.347 ± 0.262 0.05± 0.01 9.77 ± 1.68 Sewn 0.272 ± 0.031 24.617 ± 2.105 13.795 ± 1.2072.731 ± 0.284 0.130 ± 0.017 15.10 ± 1.29  laminate Errors representstandard errors from five experiments.

Example 5 Distribution of Collagen IV and Laminin in a DisruptedBasement Membrane

FM scaffold obtained according to the process set forth in Example 1, aswell as the epithelium removed during delamination of the forestomachtissue, were fixed by immersion in 7% neutral-buffered formalin andembedded in paraffin wax. Sections were cut at 10 μm thickness using amicrotome before the section was relaxed in heated water and mounted onAPE-coated slides. Sections were fixed to the slides by immersion inparaformaldehyde for 10 minutes and stored in dust-free conditions atroom temperature. Paraffin from slides was dissolved by immersion infour 5 minutes washes of 100% xylene and rehydrated through descendingconcentrations of ethanol before immersion in 50 mM TBS (pH 7.4).Endogenous peroxidases were quenched using 5% H₂O₂ in 70% methanol for30 minutes. Staining procedures were carried out according to theprotocols of a DAB secondary detection kit (Chemicon). Sections werewashed three times in TBS for 5 minutes each time, followed byincubation in blocking serum for 30 minutes to block non-specificbinding. Subsequently, sections were incubated in an appropriate primaryantibody (i.e., an antibody which recognizes laminin or collagen IV,such as a rabbit anti-ovine primary antibody), at a dilution of 1:100(both primary antibodies were optimized in the tissue prior to use inthis assay). Sections were incubated in primary antibody in a humidifiedchamber for 30 minutes at ambient temperature. Sections were then washedin TBS containing 0.1% Triton X-100 (TrTBS), and then twice in TBS for 5minutes each. Sections were incubated with the secondary antibodyprovided in the DAB secondary detection kit, described above, for 10minutes at room temperature. Sections were subsequently washed once inTrTBS and twice in TBS followed by incubation with strepavidin-biotinconjugate (Vector Laboratories, USA) at room temperature for 10 minutes.Slides were washed three times in TBS prior to incubation with thechromogen diaminobenzidine tetrachloride (DAB) for 1-5 minutes to allowmaximal antigen staining with minimal background staining. DAB labelingwas halted by washing sections in dH₂O. Sections were thencounterstained with haemotoxylin, rewashed in running tap water,dehydrated in ascending concentrations of ethanol, cleared in xylene,mounted using DPX mountant and stored at room temperature until viewing.Labeled proteins appeared brown, whereas the haemotoxylin-labeled nucleiappeared blue. Negative controls were carried out by omitting theprimary and/or secondary antibody application during the aboveprocedures. Slides were viewed under a light microscope and photos weretaken using AnalySIS software.

Immunohistochemistry of the FM scaffold revealed collagen IV and lamininwere localized to the epithelial and vascular basement membranes andwere also present within the dense layer of matrix deep within thelamina propria. Laminin and collagen IV staining of the FM scaffoldrevealed that the basement membrane was not a continuous surface, butrather was discontinuous or disrupted. Collagen IV and laminin wereadditionally seen in samples of epithelium taken after delamination ofthe tissue. Staining of the epithelial tissue revealed that collagen IVand laminin were localized to fragments of basement membrane. Thepresence of laminin and collagen IV on both the epithelial layer and theluminal surface of the FM scaffold indicates that the basement membraneis disrupted during delamination of the forestomach tissue. Disruptionand fracture of the basement membrane during processing leads to releaseof the epithelium. Significantly, the presence of laminin and collagenIV in the lamina-propria layer of the FM scaffold provides a source ofthese important cell adhesion molecules during tissue regeneration.

Example 6 Basement Membrane Disruption and Fracture

Suspending an ovine forestomach containing NaCl in a hypotonicprocessing solution which contains 0.028% Triton X-200 and 0.1% EDTA for16 hours resulted in shedding of the epithelium from the underlyingpropria submucosa as the basement membrane was disrupted.

Analysis by immunohistochemistry of sections taken from the epithelialsheets showed the presence of collagen IV and laminin in the remnants ofthe basement membrane on both the FM and the shed epithelial layer(described above). Based on the above, it is clear that the processingsolutions are disrupting the structure of the basement membrane, causingthe basement membrane to be fracture and the epithelial layer to bereleased. Following processing, the epithelium peeled away in sheetsfrom the underlying propria-submucosa. This “sheeting” of the epitheliumoccurred following immersion in Triton X-200 and EDTA solution duringthe STOF process.

Western blotting revealed that laminin, a major component of thebasement membrane, was being released into solution during the STOFprocess. This was evident when solutions of either 0.028% Triton X-200,or 0.028% Triton X-200+0.1% EDTA, were sampled during the STOF process.Proteins present in the samples were separated by gel electrophoresisand visualized using an anti-laminin antibody (FIG. 8). Laminin wassolubilized by either of the 0.028% Triton X-200 containing solutions(Lanes 2 and 4, FIG. 8) but not by a solution containing NaCl (Lane 3,FIG. 8) or 0.1% SDS (Lane 5, FIG. 8).

A biochemical analysis of the FM scaffold and the epithelial layer shedduring processing revealed that laminin, a principle basement component,was present in both fractions. Laminin in both the FM scaffold and theepithelial tissue were quantified using ELISA according to Example 7,below. Laminin concentrations in the FM scaffold and epithelial tissuewere 5.87±2.16 and 17.3±1.1 μg/g, respectively. The fact that lamininwas detected in relatively high concentrations in the shed epithelialtissue further supports the observation that during STOF processing thebasement membrane of the forestomach tissue was fractured, leading toloss of the epithelial layer.

Example 7 Biochemical Composition of FM Scaffold

An extensive study was undertaken to understand both the major and minorcomponents of FM scaffold, e.g., scaffold obtained according to theprocess set forth in Example 1. Macroscopically the product can beconsidered as a collagen matrix that will support cellular infill anddifferentiation. However, the manufacturing process has been developedin such a way as to retain minor biologically active components, forexample growth factors and glycosaminoglycans (GAGs). These minorcomponents play an equally important role in wound healing and theirpresence in FM scaffolds imparts beneficial wound healing properties tothe product.

In all cases the biochemical composition of the FM scaffold was comparedwith the porcine small intestine submuscosa (SIS), and the raw material,ovine forestomach.

In all cases, tissue samples were firstly frozen in liquid nitrogen andmilled in a spice grinder to yield fine particulate. The powder wasextracted and analyzed for biochemical macromolecules according toestablished procedures. Biochemical analysis of the major components issummarized in Table 5.

Total soluble collagen was quantified by an enzymatic digestion of thepowdered samples (5 mg/mL pepsin, 0.5 M acetic acid, 37° C., 16 h),followed by centrifugation and analysis of the supernatant using theSircol™ Soluble Collagen Kit (Biocolor—County Antrim, UK). Total solublecollagen of the samples was calculated from a standard curve of rat tailcollagen I (Gibco Invitrogen—California, USA). The concentration oftotal soluble collagen was approximately equal in the ovine forestomach,FM scaffold and SIS.

While soluble collagen quantification may represent the relativeabundance of physiologically available collagen, it does not take intoaccount the pepsin-insoluble collagen components of the matrices. Thus,to understand total collagen content of the test samples, ahydroxyproline analysis was conducted according to establishedprocedures. Samples were hydrolyzed in 6 M HCl (120° C., 60 minutes),then the hydroxyproline residues reacted to form a pyrrole chromophorethat was quantified using absorbance. Total collagen was calculated froma standard curve of hydroxyproline (Sigma—Missouri, USA), where it wasassumed the ratio of hydroxyproline to total collagen is 7.14. In the FMand SIS scaffolds, collagen was approximately 80-80% by composition. Asexpected, pepsin-soluble collagen represented only a small fraction ofthe total collagen present in the samples.

Collagen IV is an important component of the basement membrane thatpromotes cell adhesion and proliferation. The basement membrane of FM ispartially intact, suggesting that this feature might impart beneficialproliferative properties to the product. Collagen IV was solubilizedwith 4% SDS (37° C., 16 hours) in extraction buffer, dialyzed againstPBS, quantified using a direct ELISA probed with anti-collagen IVantibody (Abacus ALS-Queensland, Australia), and detected with asecondary antibody conjugated to HRP (Abacus ALS-Queensland, Australia).Collagen IV was quantified relative to a standard curve of partiallypurified bovine collagen IV (Chemicon Millipore—Massachusetts, USA).Collagen IV represented a small proportion of total collagen in thematrices (approximately 2%).

Elastin is an important structural protein which forms a network ofelastic fibers within the ECM to provide resilience, texture, durabilityand the ability to recoil after stretching. Elasticity of the protein isimparted by extensive cross-linking of the soluble tropoelastin monomersto yield an extensive covalent array. Given its extensive lysinecrosslinking, elastin is especially insoluble and therefore difficult toquantify in biological samples. In order to understand the relativelevels of elastin, both ‘soluble’ and ‘insoluble’ elastin werequantified. Samples of matrices were solubilized in 0.25 M oxalic acid(105° C., 16 hours), centrifuged to pellet out the insoluble material,then the supernatant analyzed using the Fastin™ Elastin Kit(Biocolor—County Antrim, UK), according to the manufacturersrecommendations. Solubilized elastin was quantified relative to astandard curve of partially purified bovine elastin. The concentrationof soluble elastin composition was approximately ten-fold less in FMrelative to SIS. Insoluble elastin was quantified using a mass balanceassay according to established procedures. Briefly, samples wereextracted with (1) ethanol/diethyl ether (1:1, 15 minutes, rt ° C.); (2)0.3% SDS (16 hours, rt ° C.); and (3) 0.1 M NaOH (15 minutes, 100° C.).Samples were centrifuged and the supernatant discarded after eachextraction step. Insoluble elastin remained after the extractionprocedure; as such, total insoluble elastin was calculated by comparingthe dry weight of the sample before and after the extractions. Totalinsoluble elastin concentrations were lower in the FM than in the SIS(3.0% and 5.3%, respectively).

Glycosaminoglycan molecules (GAGs), including heparan sulphate andchondroitin sulphate, bind growth factors and cytokines, and controlwater retention and gel properties within the ECM. The heparan bindingproperties of numerous cell surface receptors and of many growth factors(e.g. FGF family, VEGF) make heparan-rich GAGs extremely desirablecomponents of scaffolds for tissue repair. Total GAGs were quantifiedusing a colorimetric dye-binding assay (Blyscan Sulfated GlycosamineKit; Biocolor—County Antrim, UK) following papain (125 μg/mL) digestionof the powdered samples. GAG concentration was calculated from astandard curve of chondroitin sulphate (Sigma—Missouri, USA). SIS hadhigher concentrations of total sulphated GAGs than both ovineforestomach and FM (7.3±0.4, 3.9±0.1 and 0.6±0.1 mg/g, respectively).Heparan sulphate concentrations were 0.2 mg/g and 2.1±0.1 mg/g for FMand SIS, respectively. A more detailed discussion of the quantificationof heparan sulphate is provided below. The concentrations of hyaluronicacid (HA) in each of the three samples were determined using an ELISAkit (Echelon Biosciences—Utah, USA). HA was found to be present in lowconcentrations in all three samples.

A critical feature in the manufacture of FM are steps that decellularizethe ECM and thereby reduce any negative host response to the xenoplant.Typically, decellularization is achieved through detergent-mediateddisruption of cell membranes, leading to cell lysis and solubilisationof cellular components. The presence of nucleic acids was used as asurrogate marker for the cellularity of the matrices. DNA fragments intheir own right also pose some risk off invoking a host-mediated immuneresponse. Total DNA was quantified using a fluorescent dye-bindingassay. Papain digested samples were incubated with Hoechst 33258 dye (10μg/mL, Sigma—Missouri, USA), and relative fluorescence units werequantified using a microtitre plate reader. Total DNA concentration wascalculated from a standard curve of calf thymus DNA (Sigma—Missouri,USA). The concentration of DNA in FM was less than that in SIS (0.2% and0.4%, respectively). As expected, there was a significant reduction inDNA in FM relative to the ovine forestomach raw material (0.2% and 2.6%,respectively).

Tissue lipids are found primarily in cell membranes and therefore offeranother surrogate that can be useful in assessing the extent ofdecellularization of the ECM matrices. The lipid content of the FMscaffold was determined using mass balance following ether extraction ofthe samples. Both FM and SIS scaffolds had approximately 6% lipidcomposition, while ovine forestomach had a lipid composition of 14.4%.

TABLE 5 Major biochemical components of the ovine forestomach, and FMand SIS tissue scaffolds Ovine Forestomach FM SIS % % % (mg/g ± SE)¹composition (mg/g ± SE)² composition (mg/g ± SE)¹ composition³ Total142.7 ± 9.7  40.5 821.0 ± 9.0  89.72 629.7 ± 39.7 80.6  Collagen Soluble 55.3 ± 3.21 —³ 51.7 ± 3.3  —³ 48.7 ± 5.3 —³ Collagen Collagen III 61.3± 0.7 —³ 196.8 ± 6.9  —³ 171.5 ± 11.1 —³ Collagen 46.6 ± 1.5 —³ 9.7 ±2.1 —³  6.6 ± 1.1 —³ IV Soluble 112.0 ± 15.0 32.0 4.8 ± 0.5 0.53 54.0 ±6.1 6.9 Elastin Insoluble 30.9 ± 6.4 8.8 27.5 ± 4.5  3.01 41.4 ± 5.0 5.3Elastin Total  3.9 ± 0.1 1.1 0.6 ± 0.1 0.07  7.3 ± 0.4 0.9 GAGs HeparanN.D. —⁵ 0.2 —⁵  2.1 ± 0.1 —⁵ Sulphate Hyaluronic  1.95 ± 0.02 0.6 0.4 ±0.1 0.05  1.58 ± 0.11 0.2 Acid DNA  9.0 ± 0.5 2.6 1.7 ± 0.5 0.19  2.7 ±0.2 0.4 Lipid⁶ 50.2 ± 2.9 14.4 59.0 ± 5.2  5.7 44.7 ± 4.0 5.7 ¹Errorsrepresent standard error from triplicate experiments. ²Errors representstandard errors from at least three independent production lots, testedin triplicate. ³Percentage composition based on total collagen from thehydroxyproline analysis only excludes collagen III, collagen IV andsoluble collagen. ⁴N.D. = Not detected. ⁵Percentage composition excludesheparan sulphate as this is included in total GAGs. ⁶As determined fromthe insoluble elastin - mass balance assay.

FM and SIS tissue scaffolds, as well as ovine forestomach, were analyzedfor minor biochemical components, fibronectin, laminin and the growthfactors VEGF, FGF2, TGFβ1, and TGFβ2. Results are summarized in Table 6.

Fibronectin is a glycoprotein that is distributed throughout the ECM andplays an important role in cell growth adhesion, migration anddifferentiation. Fibronectin binds collagens and heparans and,importantly, provides ligands for the adhesion of cell surface integrinreceptors leading to cell attachment and proliferation. Fibronectin wasquantified using QuantiMatrix Human Fibronectin ELISA Kit (ChemiconMillipore—Massachusetts, USA) following dialysis of samples extractedwith 4% SDS. The concentration of fibronectin in FM was significantlyhigher than SIS, 13.67±1.64 and 5.00±0.05 μg/g, respectively.

Laminin is an ECM protein which is capable of binding to type IVcollagen molecules, heparan sulphate, and integrin receptors, thusforming important connections between cells and the basement membrane orthe ECM. Laminin was quantified in samples extracted with 4% SDS using aQuantiMatrix human Laminin ELISA Kit (Chemicon Millipore—Massachusetts,USA), and concentrations were determined relative to a standard of humanlaminin. The concentrations of laminin in FM and SIS were approximatelyequal.

Basic Fibroblast Growth Factor (FGF2) is multifunctional and plays animportant role in wound healing including the promotion of endothelialcell differentiation during angiogenesis, and cell differentiation andmigration of a number of cell types. Samples were analyzed for thepresence of FGF2 using the Human FGF-Basic ELISA Development Kit(Peprotech—New Jersey, USA). Scaffold samples were extracted with 4% SDSextraction buffer and dialyzed against PBS. FGF2 was then quantifiedusing human FGF2 as a standard. Concentrations of FGF2 in the ovineforestomach and in the FM were less than the concentration shown in SIS(1.70±1.38, 0.74±0.0.09 and 4.85±0.84, respectively).

The growth factors VEGF, TGFβ1, and TGFβ2 were quantified usingcommercially available ELISA kits, according to the manufacturer'sinstructions (Bender Medsystems and Peprotech). Firstly, powdered FM wasextracted using either 2 M urea (37° C., 48 hours), or 4M guandiniumhydrochloride (37° C., 48 hours). Samples were centrifuged and thesupernatant was filtered prior to ELISA quantification. Growth factorsTGFβ2 and VEGF were also quantified using Western blot according toestablished procedures. Antibodies directed against the growth factorsand positive controls (where available) were used accordingly; theseincluded rabbit anti-TGFB2 polyclonal (Abcam®), TGFB2 monoclonal(Invitrogen™), and rabbit polyclonal anti-VEGF (Abcam®). Additionally,TGFβ1 was quantified using dot-blot employing rabbit polyclonalanti-TGFβ1 (Abcam®) and purified TGFβ1 (Invitrogen™) as a positivecontrol.

TABLE 6 Minor biochemical components of the ovine forestomach, and theFM and SIS scaffolds ¹Ovine Forestomach ²FM ¹SIS (μg/g ± SE) (μg/g ± SE)(μg/g ± SE) Fibronectin 15.30 ± 1.17  13.67 ± 1.64  5.00 ± 0.50 Laminin6.30 ± 0.24 5.87 ± 2.16 6.00 ± 0.30 FGF2 1.70 ± 1.38 0.74 ± 0.09 4.85 ±0.84 TGFβ1 N.T 0.19 ± 0.01 N.T TGFβ2 N.T 0.02 ± 0.01 N.T VEGF N.T 0.09 ±0.03 N.T ¹Errors represent standard error from triplicate experiments.²Errors represent standard errors from three independent productionlots, tested in triplicate. N.T. = not tested.

Example 8 FM Scaffold Contains the Glycosaminoglycan (GAG) HeparanSulfate

The N-sulphated GAG heparan sulphate is an important ECM-bound GAG thatplays an important role as a co-factor to FGF2 bioactivity. Heparansulphate is required for the bioactivity of FGF2 as it binds directly toFGF2 receptors in the presence of FGF2, thus stabilizing theFGF2-receptor complex. Heparan sulphate also binds free FGF2,stabilizing the growth factor and prolonging its circulating half-life.According to U.S. Pat. No. 4,902,508 and U.S. Pat. No. 6,099,567, smallintestinal submucosa contains heparan sulphate whereas stomach submucosadoes not. The absence of heparan sulphate in stomach submucosa limitsany bioactivity associated with FGF2 in ECM scaffolds derived fromstomach submucosa.

Forestomach submucosa was analyzed to determine the presence of heparansulphate. Papain digested samples of FM were resolved by celluloseacetate GAG gel electrophoresis according to establshed procedures. GAGspresent in the FM sample migrated similarly to a standard sample ofheparan sulphate, but not chondroitin sulphate B or hyaluronic acid. Byanalyzing the densitiometry of the Alcian blue stained gel it waspossible to quantify the amount of heparan sulphate present in the FMsample. Heparan sulphate concentration was approximately 0.2 mg/g.

Heparan sulphate levels were also determined using more quantitativemethods. Total GAGs were quantified in FM using the commercial BlyscanGAG detection kit, as described above. Using this approach it waspossible to determine the concentration of total GAGs (both N-sulphatedand O-sulphated GAGs) in a sample, including chondroitin sulphates (4-and 6-sulfated), keratan sulphates (alkali sensitive and resistantforms), dermatan sulphate (containing iduronic & glucuronic acid) andheparan sulphates (including heparans). With a modification to theprocedure it is possible to cleave N-sulphated heparan sulphate polymersto their constituent monomers in the presence of O-sulphated GAGs usingnitrous acid treatment. In this way it is possible to quantitate heparansulphate as a percentage of the total GAGs present in a sample. Usingthis approach it was shown that SIS had a concentration of heparansulphate of 2.1±0.1 mg/g. Under identical conditions, surprisinglyheparan sulphate could not be detected in either FM or ovine forestomachtissue.

It is possible that the nitrous acid modification to the Blyscan assaywas ineffective at resolving heparan sulfate in FM extracts. As such,alternate methods were explored to further verify the presence ofheparan sulphate in the FM matrix.

The presence of heparan sulfate was confirmed by a heparanase digestionof an FM extract prior to total GAG analysis using the Blyscan assay. Apapain treated FM extract was digested to consitutent disaccharidesusing heparan lyases I (0.5 mU), II and III (both at 0.5 mU) (SeikagakuCorporation, Japan) at 37° C. for 24 hours. After 24 hours, anadditional lyase digestion was performed to ensure complete digestion ofthe sample. Samples before and after lyase digestion were analyzed fortotal GAGs using the Blyscan assay. Disaccharides are unreactive to theBlyscan GAG assay. Results are summarized in Table 7.

TABLE 7 Heparanase digestion of FM extracts. Concentration of lyase FMsensitive Lyase Heparan FM extract extract heparan treatment sulfate (50μg/mL) (μg/mL) (mg/g) sulfate Untreated 52 70 0.7 NA 24 hours 15 42 0.40.3 48 hours 13 46 0.4 0.2

Heparan lyase digestion of the sample significantly reduced theconcentration of total GAGs, as determined using the Blyscan assay(Table 7). For example, the concentration of heparan sulfate standards(prepared at 50 μg/mL) before and after lyase digestion were 52 μg/mLand 15 μg/mL, respectively. There was no significant difference insamples treated for 24 or 48 hours, suggesting that lyase digestion wasessentially complete after the first 24 hour incubation. Lyase digestionof a heparan sulphate standard reduced the reactivity of the sample tothe Blyscan reagent, but not to background levels. This may be explainedby the presence of additional GAGs in the heparan standard that are notsensitive to heparanase digestion, and/or that heparanase digestion ofthe standard is not 100% efficient at converting heparans to theirconstituent disaccarides. Heparan lyase digestion of the FM extractssignificantly reduced the total GAGs present in the digested samples(e.g. 70 μg/mL, 42 μg/mL before and after lyase digestion,respectively). Based on this analysis, lyase sensitive heparan sulphaterepresents approximately 40% of total GAGs present in FM. Thisrepresents a heparan sulphate concentration of 0.2 mg/g, assuming totalGAG concentration prior to lyase digestion is 0.7 mg/g. This finding isin line with the gel electrophoresis analysis described above, whereheparan sulphate concentrations were determined to be 0.2 mg/g.

FM extracts were further analyzed by HPLC to establish the presence ofchrondroitin sulphate, another major GAG that may be expected to bepresent in the FM matrix. A papain-digested extract of FM was digested(20 hours at 37° C.) with 5 U/mL chondroitinase ABC (SeikagakuCorporation, Japan) in Tris buffer (50 mM pH 8.0, 0.4 M sodium acetate,0.1% BSA) to hydrolyse the chondroitin polymer to consistuent monomers,i.e. non-sulphated chondroitin, chondroitin 6-sulphate, chondroitin4-sulphate, chondroitin 2,6-sulphate, chondroitin 2,4-sulphate, andchondroitin 2,6-sulphate. The chondroitinase digested sample was alsoanalyzed by HPLC, whereby the concentration of chondroitin monomers inthe extract was used to infer the concentration of chondroitin sulphateprior to chondroitinase digestion. Samples were centrifuged and thesupernatant analyzed by RP-HPLC. Injections (20 μL) were made to aPhenosphere™ SAX 5 um column (Phenomenex—California, USA) at 22° C. Themixture was resolved using an aqueous HCl (pH 3.5)/1.5 M NaCl in HCl (pH3.5) gradient, at 1.0 mL/min flow rate. Peaks were detected at 232 nm.

Using this HPLC method none of the expected monomers of chondrotinsulphate were detected, suggesting that chondrotin sulphate is not amajor component of the total GAGs detected in FM.

Taken together these findings imply that heparan sulphate is present inthe FM scaffold, but at lower concentrations than that found in SIS(Table 8). It is interesting to note that the total GAG concentration ofFM was determined at 0.6±0.1 mg/g, suggesting that other GAG components,excluding chondroitin sulfate, may be present in the extract that werenot resolved by gel electrophoresis.

TABLE 8 Quantification of heparan sulphate using nitrous acid hydrolysisand 1,9-dimethy-methylene blue detection. Ovine Forestomach FM SISHeparan N.T. 0.2¹ 2.1 ± 0.1^(2,3) Sulphate (mg/g) ¹As determined fromgel electrophoresis analysis, and Blyscan assay of lyase pre-treatedextracts. ²As determined from Blyscan assay of nitrous acid pre-treatedextracts. ³Errors represent standard error from triplicate experiments.N.T. = not tested.

Example 9 FM Tissue Bimodal Scaffold Structure

Ovine FM was prepared using the method outlined in Example 1. Ovineglandular stomach submucosa was also prepared from glandular stomach bydelaminating the muscle and epithelium from the submucosa and thensoaking the scaffold in water for two hours to lyse the cells.Hematoxalin & Eosin (H&E) stained slides were prepared for histologyusing standard techniques.

(i) Gross Appearance: The glandular stomach submucosa had a similarmacroscopic appearance on both the luminal and abluminal surfaces. Incontrast, the FM scaffold had visible differences in the surfacecontours between the luminal and abluminal sides of the matrix. Thepapillae of the luminal side showed markedly similar topology to therete ridges of normal skin, whereas the abluminal side, left behind whenthe tunica muscularis is removed, was smooth. The bimodal nature of thisscaffold is important in terms of its interactions with different celltypes in a healing situation.

(ii) Histology: The propria-submucosa layer is unique to the forestomachof ruminants and not present in other gastrointestinal tissue. Thelamina propria of the glandular stomach and small intestine is a looseareolar layer between the glands of the mucosa which is predominantlyremoved during the delamination process. A layer of tissue called thelamina muscularis mucosa separates the lamina propria and submucosa inthe small intestine and glandular stomach. The lamina muscularis mucosais absent in the rumen of the forestomach, and consequently the laminapropria and submucosa blend to form the propria-submucosa. The FMscaffold consisted of remnants of the basement membrane and thepropria-submucosa. Analysis of the scaffold by microscopy revealed thatthe FM had a dense layer of ECM within the lamina propria of thepropria-submucosa which accounted for approximately the top 20% of thematrix thickness. The abluminal side of the FM scaffold had a more openreticular structure. The FM scaffold had a contoured luminal surface,and a dense lamina propria on the luminal side, whereas on the abluminalside, the ECM of the submucosa was more open and reticular. Accordingly,FM scaffolds have a bimodal structure. This structure makes the FM wellsuited to encouraging epithelial regeneration on the dense luminal sideof the matrix, and fibroblast invasion on the less dense abluminal sideof the matrix when used as a medical device for tissue regeneration.

(iii) Scanning Electron Microscopy (SEM): Scanning electron micrographsof FM and glandular stomach submucosa were performed on lyophilizedtissues from a one week old calf and a 6 month old lamb to compare theluminal and abluminal surfaces. FM scaffold from lamb was prepared usingthe method set forth herein. “Unprocessed” forestomach propria-submucosawas prepared by removing the epithelial and muscle layers, but notundertaking the STOF process on the tissue. Glandular stomach submucosawas prepared from the glandular stomach of a lamb by delaminating themuscle and epithelium from the submucosa and then soaking the scaffoldin water for two hours to lyse the cells. Comparison of SEM images ofneonatal bovine FM, ovine FM, and glandular stomach clearly indicatedthat FM has two distinct surfaces and has clear sidedness, whereasglandular stomach submucosa is very similar on both sides.

Cross-sectional SEM images of FM and glandular stomach submucosademonstrated the presence of a thick, dense layer of ECM on the luminalsurface of the FM propria-submucosa, compared with the thinner and moreuniform structure of the glandular stomach submucosa. Notably, the denselayer of ECM is absent from the glandular stomach submucosa.

Example 10 Surface Area of FM Tissue Matrix

Six forestomachs and six glandular stomachs from six month old lambswere collected from a local abattoir. Grossly, the forestomach is asubstantially larger organ than the glandular stomach, and therefore isideally suited for producing large surface area scaffolds.

To compare the difference in size, the respective volumes of theforestomach and the glandular stomach were measured. It was found thatforestomachs were typically 12-15 liters in volume, whereas glandularstomachs were limited to 2.5-3 liters. Based on an approximation to thesurface area of a sphere, this represents an area of approximately 405cm² for the forestomach, compared with 104 cm² for the glandularstomach.

Table 9 summarizes the differences between ECM scaffolds derived fromovine forestomach and glandular stomach. The typical dimensions of theseECM sheets are shown in Table 9 to illustrate the approximate differencein the dimensions of scaffolds obtained from the forestomach andglandular stomach. It was also noted that the glandular submucosa matrixhad a fragile, delicate structure and was difficult to isolate, whereasthe FM was much more robust and easier to obtain.

TABLE 9 Comparison of surface area of ovine forestomachs and glandularstomachs Forestomach Glandular Stomach Volume (mL) 13,000 3,000 SurfaceArea (cm²) 2678 1014 Typical ECM sheet 15 8 width (cm) Typical ECM sheet27 17 length (cm) Area of typical 405 104 ECM sheet (cm²) Comments Easyto separate tissue Difficult to separate tissue layers to producelayers, a thin and friable FM, thick and ECM, which is difficult torobust ECM produce large intact sheets

A significant advantage of FM over other known compositions is thatlarge constructs can be produced from a single organ (e.g., theforestomach). For example, for an animal of any age or bodyweight,sheets of FM produced from the forestomach were typically 3-4 timeslarger than those produced from the glandular stomach, which in turn aretypically larger than those which can be produced from the bladder. ECMsheets obtained from tubular organs such as small intestine are limitedin width to the circumference of the organ. For example, smallintestinal submucosa from market weight pigs is limited to widths ofless than 10 cm due the circumference of the small intestine.

Example 11 Thickness of FM Scaffold

Four sheets of FM scaffold were prepared from the forestomachs of four5-6 month old lambs weighing approximately 25 kg. Thirteen adequatelyspaced measurements of the thickness of the FM sheet were made in eachof the four samples. The results are presented in the Table 10.

TABLE 10 Thickness of FM sheets derived from four 5-6 month old lambsSampled Measurements (μm) Sheet Min Max Mean Number 1 252 375 309 13 2205 268 230 13 3 270 452 364 13 4 299 463 388 13

Small intestinal submucosa derived from pigs weighing greater than 180kg (as indicated in U.S. Pat. No. 5,372,821) is typically only 100 μmthick. The above measurements demonstrate that immature lambs which areless than 20% of the size of a mature pig can provide ECM sheets whichare at least three times as thick as that obtained from small intestinalsubmucosa of a pig. Such thicker scaffolds possess advantageousproperties, such as greater strength and longer persistence followingimplantation in vivo. Thicker FM scaffolds can be produced withincreasing age and weight of the animal.

Example 12 Biaxial Strength of FM Scaffold

Samples of 1-ply FM scaffold sheets were laminated using a layer ofcollagen polymer, as described in Example 3, to give 2-, 3-, 4- and8-ply laminates. Additionally, forestomach propria-submucosa andglandular stomach submucosa were manually isolated from the forestomachand the glandular stomach, respectively. These ‘unprocessed’ sampleswere not exposed to the STOF process described in Example 1.

Ball burst strength provides a measure of a biomaterials resistance to aload when biaxial force is applied. The test is conducted by clampingthe test material in a circular orifice and forcing a metal spherethrough the centre of the sample until the sample fails, allowing thesphere to pass through. The relative strengths of materials are comparedusing the force at the point of failure of the material, termed the‘maximum compression load’ (Newtons, N). Maximum compression load of asample will be dependent on the elasticity and strength of the sample,as well as the sample thickness. The ball-burst test determines biaxialstrength, whereby forces are equally applied in all directions. Incomparison, uniaxial strength (see below) determines load to failure inone direction only.

In line with the Standard Test Method for Bursting Strength of KnittedGoods, Constant-Rate-of-Traverse (CRT) Ball-Burst Test (ASTM D 3797-89),a 24.5 mm polished steel hemisphere was pushed against the ECM sheetsuntil failure on an Instron 1122 Machine. The maximum compression loadwas defined as the force required to rupture the sheet. At least six(n=6) samples of each ECM were tested. Results are shown in Table 11 andFIG. 9. FM scaffold was as strong as unprocessed forestomachpropria-submucosa (92.8±12.7 and 114.3±8.1 N, respectively) (Table 11and FIG. 9), and both forestomach ECM materials were substantiallystronger than those derived from intestinal tissue, and glandularstomach tissue.

TABLE 11 Comparison of ball burst strength of FM scaffold and other ECMscaffolds Maximum Compression Load Scaffold (N) Ovine FM Scaffold  92.8± 12.7 Porcine Intestinal Submucosa^(c) 20.1 ± 0.5 Ovine UnprocessedStomach Submucosa 20.8 ± 1.7 Ovine Unprocessed Forestomach Propria-114.3 ± 8.1  Submucosa Porcine EtO-sterilized Urinary Bladder Matrix^(a)42.2 ± 7.7 Porcine Non-sterilized Urinary Bladder 57.3 ± 8.9 Matrix^(b)Porcine e-beam sterilized Urinary Bladder 11.3 ± 1.9 Matrix PorcineUrinary Bladder Submucosa^(c) 15.4 ± 0.5 ^(a)Freytes et al., “Effect ofStorage Upon Material Properties of Lyophilized Porcine ExtracellularMatrix Derived from the Urinary Bladder” (2005) J. Biomed. Mater. Res.B: Appl. Biomater. ^(b)Freytes et al., “Uniaxial and Biaxial Propertiesof Terminally Sterilized Porcine Urinary Bladder Matrix Scaffolds”(2007) J. Biomed. Mater. Res. B: Appl. Biomater. ^(c)U.S. Pat. No.6,099,567

A dramatic increase in the strength of the FM laminates was observed asadditional sheets were laminated to generate a series of multi-plydevices (2-, 3-, and 4-ply). For example, 4-ply ovine FM had a maximumcompression load of 361.5±24.9 N, while 1-ply FM had a maximumcompression load of (92.8112.7 N), as shown in FIG. 10.

The biaxial strength of a 4-ply FM laminate was compared with publishedball burst data for the commercial ECM-based products, Alloderm™(LifeCell Corporation), Strattice™ (LifeCell Corporation), and Surgisis™(Cook Biotechnology) (see Table 12). The 4-ply FM had a lower maximumcompression strength than the other implant products. However, incomparing ball-burst strengths it is important to consider also thethickness of the test material, as this dimension will significantlyimpact the observed maximum compression load. For example, the reportedthickness of Alloderm™ was 1.9±0.13 mm, while the 4-ply FM product wasapproximately 75% thinner at 0.47±0.01 mm. To take into account thedifferent thickness of the ECM products and therefore make a meaningfulcomparison of the biaxial strengths of the ECM products, the maximumcompression load (N) of the products was normalized to their thicknesses(mm). Using this analysis, the relative biaxial strengths of the fourproducts were found to be statistically similar (see Table 12 and FIG.11).

TABLE 12 Comparison of ball-burst properties of 4-ply FM and commercialECM-based implant products Normalized Maximum Maximum CompressionThickness Compression Load Load (N ± SE) (mm) (N/mm ± SE) 4-ply FMScaffold 361.5 ± 24.9 0.47 ± 0.01 773.7 ± 68.6  Alloderm ™ 1781.5 ±80.2  1.9 ± 0.1   937 ± 106.0 (Boguszewski, Dyment et al. 2008)Strattice ™ 1059.7 ± 181.8 1.49 ± 0.07 711.2 ± 155.4 (Boguszewski,Dyment et al. 2008) Surgisis ™ (package 440 ± 81 0.76 578.9 insert)Errors represent standard errors from at least five samples, or frompublished data. No thickness error reported for Surgisis ™.

Example 13 Uniaxial Tensile Strength

Uniaxial strength measures the one-dimensional force tolerance of abiomaterial whereby a strip of material is clamped at either end andopposing forces are applied. The force at failure of the material istermed the ‘maximum load’ (N).

The maximum load of a sample is dependent on the inherent strength ofthe test material, as well as the size and thickness of the test sample.Uniaxial testing was performed on an Instron 1122 Machine. Samples werecut to a dog-bone shape with a mid-substance width of 1 cm, aspreviously described (see Freytes et al., “Effect of Storage UponMaterial Properties of Lyophilized Porcine Extracellular Matrix Derivedfrom the Urinary Bladder” (2005) J. Biomed. Mater. Res. B: Appl.Biomater. and Freytes et al., “Uniaxial and Biaxial Properties ofTerminally Sterilized Porcine Urinary Bladder Matrix Scaffolds” (2007)J. Biomed. Mater. Res. B: Appl. Biomater., both incorporated herein byreference in their entirety).

FM scaffold was prepared according to Example 1. Additionally,‘unprocessed’ forestomach propria-submucosa and stomach submucosa wereprepared manually. These unprocessed samples were not exposed to theSTOF process. All samples were mounted to the Instron test apparatus andpulled to failure at a constant rate of 20 mm/min. At least eight (n=8)samples of each ECM were tested.

The results, as shown in Table 13 and FIG. 12, demonstrated thatunprocessed forestomach propria submucosa is much stronger than stomachsubmucosa from the same animal. Additionally, ovine FM scaffold from sixmonth animals (approx. 25 kg) was stronger than porcine stomachsubmucosa derived from mature animals (approx. 50 kg), as reported inthe U.S. Pat. No. 6,099,567. This data indicates that FM scaffold ismuch stronger than stomach submucosa, irrespective of the age, andtherefore the bodyweight, of the animal. Accordingly, FM is less likelyto suffer damage or mechanical failure when used as a single sheet intissue reinforcement applications relative to stomach-derived ECM. Also,when it is desirable to achieve a composition of high tensile strengthby laminating sheets of ECM together it will be appreciated that fewerFM sheets are required compared to other gastrointestinal or urogenitalderived ECM scaffolds, making this process simpler and more costeffective.

The inherent strength of FM also makes the use of FM isolated from fetalor neonatal tissue practical, whereas other fetal or neonatal submucosaltissue sources yield tissue which is very weak due to its thinness andimmature state.

TABLE 13 Comparison of uniaxial strength of ECM scaffolds Maximum Load(N) Ovine FM Scaffold 31.6 ± 2.7  Ovine Unprocessed Stomach Submucosa2.5 ± 0.4 Ovine Unprocessed Forestomach Propria-Submucosa 17.9 ± 1.2 Porcine EtO-sterilized Urinary Bladder Matrix^(a) 4.7 ± 0.8 PorcineNon-sterilized Urinary Bladder Matrix^(b) 5.4 ± 0.5 Porcine e-beamsterilized Urinary Bladder Matrix^(a) 1.8 ± 0.2 ^(a)Freytes et al.,“Effect of Storage Upon Material Properties of Lyophilized PorcineExtracellular Matrix Derived from the Urinary Bladder” (2005) J. Biomed.Mater. Res. B: Appl. Biomater. ^(b)Freytes et al., “Uniaxial and BiaxialProperties of Terminally Sterilized Porcine Urinary Bladder MatrixScaffolds” (2007) J. Biomed. Mater. Res. B: Appl. Biomater.

In addition to the foregoing data, the maximum uniaxial load for singleand multi-ply FM scaffolds was determined, as well as the maximumtangential stiffness (N/mm), elongation at failure (mm), the modulus ofelasticity (GPa) and yield stress (MPa). Yield stress is a termnormalized to the dimensions of the test sample and can be used tocompare similar products of different thicknesses. This term is ameasure of the inherent strength of the material. The modulus ofelasticity, or Young's modulus is a measure of the elastic potential ofthe material, that is its ability to be deformed without failure andreturn to its original state. The modulus of elasticity is an intrinsicproperty of the material, thus allowing samples of different sizes to becompared. For the modulus of elasticity, a lower number indicatesgreater elasticity. For example, the modulus of rubber is 0.01-0.1 GPa.

A comparison of the uniaxial strength properties of 1-ply through to4-ply FM laminates is presented in FIG. 13. As expected, increasing thelamination state and hence thickness of the product significantlyimproved uniaxial strength and stiffiness. However, lamination did notstatistically alter the maximum elongation, or modulus of elasticity.This indicates that the process of lamination increases the strength ofthe product, but does not alter its pliability, suggesting the handlingof 1- and multi-ply products would be similar.

In order to evaluate the relative uniaxial strength of ovine FMproducts, a literature survey was performed to identify and extractrelevant data relating to the strength of alternate ECM-like products.The uniaxial strength properties and thicknesses of the 1- and 2-ply FMscaffolds were compared with published data for the dural repairproducts DuraGuard™ and Durarepair™ (Table 14 and FIG. 14). The 1- and2-ply FM scaffolds had statistically equivalent yield stress (10.15±1.81and 9.77±1.68 MPa, respectively). This is expected as yield stress isnormalized to the sample thickness, such that two samples prepared fromthe same material but of different thicknesses should give identicalyield stress. The 1- and 2-ply FM products had a similar yield stress toDura-Guard™ (13.5±3.34 MPa), and all three products out performedDuraDerm™ (6.27±4.20 MPa). Durarepair, the strongest of the products,had a yield stress of 22.7±2.83 MPa. The 1- and 2-ply FM scaffolds(Young's modulus of 0.04+0.01 and 0.05±0.01 GPa, respectively) hadbetter elastic properties than Dura-Guard™ and Durarepair™ (0.08+0.02and 0.07±0.01 MPa, respectively). DuraDerm™ demonstrated the bestelastic potential (modulus 0.002±0.009 GPa), but as noted above, was theweakest of the products surveyed.

This analysis suggests that FM scaffolds have strength and elasticcharacteristics that are equivalent to or better than current commercialdural repair products. This analysis indicates that a 4-ply FM scaffoldhas an equivalent strength to commercial dural products.

TABLE 14 Comparison of uniaxial strength properties of 1-ply and 2-plyFM scaffolds with commercial dural repair products Modulus of ElasticityYield (Youngs') Stress Thickness (GPa ± SE) (MPa) (mm ± SE) 1-ply FMScaffold 0.04 ± 0.01 10.15 ± 1.81 0.25 ± 0.01 2-ply FM Scaffold 0.05 ±0.01  9.77 ± 1.68 0.31 ± 0.01 DuraDerm ™ (Sclafani, 0.002 ± 0.009  6.27± 4.20 1.4 ± 0.2 McCormick et al. 2002) Dura-Guard ™ (Zerris, 0.08 ±0.02 13.5 ± 3.3 0.400 ± 0.001 James et al. 2007) Durarepair ™ (Zerris,0.07 ± 0.01 22.7 ± 2.8 0.50 ± 0.02 James et al. 2007) Errors representstandard errors from at least five samples, or published values.

To evaluate the suitability of laminated FM scaffolds for implantation,biophysical properties of the scaffolds, as derived from uniaxialstrength testing, were compared with similar published data relating tothe commercially available product Alloderm™ (see Table 15). It isapparent from the spread of reported data that a consensus about thetrue strength and elastic potential of Alloderm™ has not been reached inthe literature. This may reflect the cadaveric origins of the product,leading to product irregularities, including the observed variability inthickness. The reported yield stress of Alloderm™ ranges from 7.00±1.00to 16.79±2.10 MPa. In order to make a meaningful comparison, themid-point of the data spread was calculated as 11.90 MPa, which iscomparable to the yield stress of the 4-ply laminate (11.97±1.16 MPa).The two reported modulus of elasticity for Alloderm™ are an order ofmagnitude different from each other, making a comparison with the 4-plyproduct difficult. However, the 4-ply product retains the elasticproperties of the 1- and 2-ply products and is similar to those of thedural repair products Dura-Guard™ and Durarepair™ (see discussionabove).

Accordingly, the studies performed herein indicate that FM scaffoldshave strength characteristics equivalent to the material used tomanufacture Alloderm™. In particular, the data indicates that multi-plyFM scaffolds have suitable strength and elastic properties for implantapplications, where inherent strength is a requirement of the implantmaterial.

TABLE 15 Comparison of uniaxial strength properties of 4-ply FM scaffoldand Alloderm ™ Modulus of Elasticity (Youngs') Yield Stress Thickness(GPa ± SE) (MPa) (mm ± SE) 4-ply FM Scaffold 0.06 ± 0.01 11.97 ± 1.160.47 ± 0.01 Alloderm ™ N.D 16.79 ± 2.10 N.D (Lemer, Chaikin et al. 1999)Alloderm ™ N.D 10.55 ± 2.37 0.9 ± 0.1 (Morgan, McIff et al. 2004)Alloderm ™ N.D  7.20 ± 2.56 1.00 ± 0.05 (Choe, Kothandapani et al. 2001)Alloderm ™ N.D 15.25 ± 7.13 1.34 ± 0.05 (Vural, McLaughlin et al. 2006)Alloderm ™  0.014 ± 0.0015  7 ± 1 0.45 ± 0.05 (Gouk, Lim et al. 2008)Alloderm ™ 0.001 ± 0.002  8.64 ± 3.31 1.89 ± 0.30 (Sclafani, McCormicket al. 2002) N.D = no data. Errors represent standard errors from atleast five samples, or published values.

Example 14 Suture Retention Strength

Suture retention testing is a practical clinical consideration thatdetermines the resistance of a biomaterial to suture ‘pull-out’. Thetest protocol is similar to uniaxial strength, however in this exampleone edge of the test material was clamped, while the other was securedto an opposing clamp via a suture placed through the test material. Thesuture was secured to the biomaterial at a defined ‘bite-depth’, that isthe distance from the site of the suture to the edge of the testmaterial. Opposing forces were applied at a constant rate until thematerial failed and the suture was pulled-out of the test material. Theload at failure of the material, termed the ‘maximum load’ (N), isdependent on the inherent strength of the test material, the bite-depthand thickness of the sample.

A series of laminated FM scaffolds were tested for suture retention. Asexpected, lamination of FM sheets increased the resistance of theproduct to suture pull-out, as shown in FIG. 15. For example, the loadto failure of the 1-ply FM was 5.91±0.60 N, while the stronger (asdetermined from uniaxial and ball-burst testing, above) 4-ply FMscaffold gave a load to failure of 15.96±1.30 N.

As seen in FIG. 15, observed suture retention strength is dependent onsample thickness. Thus, in order to compare the suture retentionstrength of ovine FM laminates with commercial products using publisheddata, the maximum load to sample thickness was normalized. A comparativeanalysis of FM scaffolds and the dural repair products Durarepair™ andDura-Guard™ is shown in Table 16 and FIG. 16. The suture retentionstrength of the 1-(4.7±0.4 N) and 2-ply (7.1±0.5 N) FM scaffolds wasless than Dura-Guard™ (10.02±1.35 N) and Durarepairm (12.38±2.10 N).However, taking into account the relative thicknesses of the fourproducts via normalized suture retention, the four products hadequivalent potential to resist suture pull-out.

TABLE 16 Comparison of the suture retention strength of FM scaffolds andcommercially available dural repair matrices Suture Retention -Normalized Maximum Thickness Suture Retention Load (N ± SE) (mm ± SE)(N/mm) 1-Ply FM Scaffold  4.7 ± 0.4 0.25 ± 0.01 18.9 ± 1.5 2-ply FMScaffold  7.1 ± 0.5 0.31 ± 0.01 23.2 ± 1.6 Dura-Guard ™ 10.02 ± 1.350.40 ± 0.01 25.1 ± 3.4 (Zerris, James et al. 2007) Durarepair ™ (Zerris,12.38 ± 2.10 0.50 ± 0.02 24.76 ± 5.19 James et al. 2007) Errorsrepresent the standard error of five samples, or published data.

The comparative study was extended to determine the performance oflaminated FM scaffolds relative to the commercially available implantproducts Alloderm™, Strattice™ and Surgisis™. The results are shown inTable 17 and FIG. 17. It is important to note that while the ASTMstandard for suture retention testing prescribes a 2 mm bite depth,various bite depths were employed across these studies, from about 2 mmto about 10 mm (Table 17). Increasing the bite depth used during testingincreases the observed maximum load as more force is required to pullthe suture further through the sample until the edge is reached andpullout is observed. The 4-ply FM laminate scaffold was significantlymore resistant to suture pull-out than the Surgisis™ product (normalizedsuture retention strengths, 34.2±2.8 and 18.0 N/mm, respectively). Thereported suture retention strengths of both Alloderm™ and Strattice™have employed a modification to the ASTM standard, using a bite-depth of10 mm (Table 17). The normalized suture retention strengths of theseproducts was 71.2±10.71 and 40.2±3.43 N/mm, respectively. In comparison,the normalized suture retention strength of the 4-ply FM scaffold was34.2±2.8 N/mm, using a bite-depth of 2 mm. This data indicates that thesuture retention strength of the 4-ply FM scaffold would likely beequivalent to both Alloderm™ and Stratticem if all three products weretested in side-by-side experiments using equivalent bite-depths. Thisnotion is supported by the observation that 4-ply FM scaffold hassimilar yield stress under uniaxial testing (see Table 15), and that thethree products have similar normalized ball-burst strengths (see Table12).

TABLE 17 Comparison of the suture retention strength of laminated FMscaffolds and commercially available implant matrices Suture NormalizedRetention - Suture Bite Maximum Thickness Retention Depth Load (N ± SE)(mm ± SE) (N/mm) (mm) 4-ply FM 16.0 ± 1.3 0.47 ± 0.01 34.2 ± 2.8  2Scaffold Alloderm ™ 135.2 ± 11.1  1.9 ± 0.13 71.2 ± 10.71 10(Boguszewski, Dyment et al. 2008) Strattice ™ 59.9 ± 2.3 1.49 ± 0.0740.2 ± 3.43  10 (Boguszewski, Dyment et al. 2008) Surgisis ™ (Oasis 13.7± 3.2 0.76 18.0 2 Product Insert) Errors represent standard errors fromat least five samples, or from published data. No error reported forSurgisis ™.

Example 15 Hydrostatic Permeability Index (PI) of FM Scaffold

The permeability of implantable ECM scaffolds can influence the rate ofcell infiltration and diffusion of molecules into and from the exogenousimplant. The desired permeability of the implant will depend on theapplication. For example, implant devices for dural replacement shouldgenerally be relatively impermeable to protect against the potentialloss of cerebral spinal fluid. Implants for tissue reconstructionrequire relatively greater permeability to allow exchange ofadvantageous growth factors and cell infiltration and release of woundexudate. Thus, ideally, tissue scaffold technologies, such as the oneprovided by the present invention, allow permeability to be tailored asrequired by the target application.

In order to assess the extent of aqueous permeability of single andmulti-ply ovine FM products, permeability indices (PI) were determinedusing a hydrostatic permeability test rig, according to establishedprocedures (Freytes, Tullius et al. 2006). Given that the FM scaffold isanisotropic, having faces derived from the epithelial junction andmuscle junction, the present study sought to determine if the 1-plyscaffold had differential permeability depending on the direction offlow through the anisotropic material. Differential permeability as aresult of the “handedness” of biologically derived ECM has beenpreviously noted. Table 18 compares the permeability of 1-ply FMscaffold with published permeability indices for ECM derived fromporcine urinary bladder matrix. The results showed that FM scaffold waspermeable to aqueous solutions. Notably, the 1-ply FM scaffold wasapproximately ten-fold less permeable than the urinary bladder matrix inboth directions.

The present study was extended to determine the aqueous permeability ofnative bovine dura and 1- and 4-ply FM scaffolds. Dura was extractedfrom an approximately 2 year old cow and the dural membrane tested forpermeability using the established procedure. The dura is considered tobe a relatively impermeable membrane as it has evolved to protect themammalian central nervous system from leakage and foreign body insult.The permeability of the dura was similar to the 1-ply FM scaffold(0.0022±0.0003 and 0.0031±0.0005 mL/cm²/min, respectively), while the4-ply FM scaffold was approximately 10-fold less permeable0.00010±0.00001 mL/cm²/min). Thus, the 1-ply scaffold provides theadvantage of having a permeability very similar to native dura. Thedifference between the 2-ply and 4-ply FM scaffolds likely reflects aproperty of the polymer used to laminate the sheets. Thus, subsequentlaminate products could be developed with ‘tailored’ permeability, afeature that would be particularly relevant to dural implant products.

TABLE 18 Permeability indices of the 1- and 4-ply FM scaffolds, comparedwith urinary bladder matrix and bovine dura Permeability Index(mL/cm²/min) luminal→abluminal abluminal→luminal ratio 1-ply FM scaffold0.0031 ± 0.0005 0.0025 ± 0.0006 1.23 urinary bladder 0.08 ± 0.03 0.02 ±0.08 4.00 matrix(Freytes, Tullius et al. 2008) 4-ply FM Scaffold 0.00010± 0.00001 N.A Bovine Dura 0.0022 ± 0.0003 N.A Errors represent standarderrors form at least five samples or from published data. N.A. = notapplicable.

Example 16 In Vivo Efficacy of FM Scaffold in a Rodent Model of WoundHealing

The following study was performed to show the efficacy of FM in woundhealing. Forestomach matrix prepared from a 1 week old calf was cut intoelliptical implants (25×12 mm). Twelve male Lewis rats aged between20-23 weeks with an average weight of 370 g were obtained. The skin ofthe rats was disinfected with 0.5% chlorhexidine and 70% ethanol. A fullthickness wound was created with a sterile 12 mm biopsy punch aftermeasuring the cranial edge to be 6 cm from the base of the skull alongthe spinal axis. The matrix was implanted so that it covered the woundand extended subcutaneously beneath the skin at the cranial margin.Sterile PBS was used to re-hydrate the matrix and the wound was coveredwith Intrasite Gel (Smith and Nephew). Six similarly wounded rats weretreated with sterile saline as a control. In the first 4 days of theexperiment the rats treated with FM had a more rapid rate of closurecompared with the controls. After this time, the healing rate of the FMwound was slower compared to the controls, presumably because thescaffold reduced the degree of wound contraction. By 23 days the FMtreated wounds and the control untreated wounds were less than 20% ofthe original size. There was no evidence of inflammatory or immunereaction to the FM in the wound or in the region of the subcutaneousimplant.

Example 17 In Vivo Efficacy of FM Scaffold in a Porcine Model of WoundHealing

In order to further evaluate the in vivo performance of an FM scaffold,and to determine its effectiveness in stimulating tissue regenerationand ability to undergo remodeling, a comprehensive wound healing studyin a porcine model system was conducted. The porcine wound healing modelis generally considered to be an accepted animal model to study thewound healing process and the effectiveness of clinical or therapeuticinterventions (Lindblad 2008). This is because healing of the porcinedermis most resembles the healing process in humans. For example, woundcontracture is the dominant mechanism of wound closure in rodents, whilein pigs and humans wound closure predominantly occurs via infill of thetissue deficit (Lindblad 2008).

It is important to note that while the current study focused on an acuteporcine model of wound healing, the FM scaffold of the present inventionmay be used for both acute and chronic indications. The study proceededas follows. On day 0, a total of 20 full thickness 20 mm diameter woundswere surgically created on the back of a 6 week old anaesthetised femalepig (approx. 18-20 kg) using a dermal punch. The wounds were created infour columns of 5 rows, spaced 3 cm apart, as shown in FIG. 18. A totalof five animals were used in the study (100 total wound sites).

Each of the wounds was either untreated, or treated with either sterile1-ply FM, 2-ply laminated FM, or the established ECM-based product, SIS.In each case a circular piece of one of the foregoing scaffolds (20 mmdiameter) was applied to the wound and rehydrated in situ by theapplication of sterile saline. In order to average any positional bias,the location of the four treatments was changed between each animal,such that no two animals received the same treatment layout. Treated anduntreated wounds were dressed identically.

On days 0, 3, 7, 14, 28 and 42 all wounds were digitally imaged and thewound area and depth (height, if appropriate) were recorded.Additionally, a single row of wounds from each animal was biopsied. Thebiopsy was surgically excised and included the wound bed as well as aportion of normal tissue from the wound margins. All biopsies wereformalin fixed, mounted, sectioned and stained for analysis. Sectionedtissues were stained with H&E, Verhoff von Gieson, and Gomori'strichrome stains. Tissues were also stained using immunohistochemistrywith markers of cell differentiation, endothelialisation and immuneresponse.

(A) The FM Scaffold is Remodeled and Infiltrated with Cells During WoundHealing

An examination of fixed tissue biopsies taken during the course of thestudy indicated that the FM scaffold was infiltrated by cells during thehealing process. ECM matrices appeared as green ribbons in Gomori'strichrome stained sections, and the ECM matrices were especiallyprominent at day 7. Cells were clearly visible within the exogenous ECMscaffold at day 7. Both the FM scaffold matrices and the commercialproduct SIS were visible for approximately 14-28 days, after which timethe matrices were fully degraded and mature collagen was laid down in aprocess of remodeling.

(B) Persistence of the FM Scaffold in Treated Wounds

The persistence of FM scaffolds in the porcine wound healing model wasevaluated. In order to qualitate the persistence of the ECM treatmentsin the healing wounds, Verhoff van Gieson (elastin) stained tissuesections were examined for the appearance of the matrix, or matrixfragments, in each. In elastin-stained sections, scaffolds (1- and 2-plyFM or SIS) appeared as a red-to-black ribbon that could clearly bedistinguished from the regenerating wound. The persistence of scaffoldsat the time points sampled is summarized in FIG. 19.

At earlier time points (days 3 and 7), scaffold was clearly visible inthe top ⅓ of the wound, typically in association with the regeneratingepithelium and/or crust. As the time course progressed, the scaffoldappeared to be migrating into the wound bed and undergoing degradation.There was little difference in the longevity of the three scaffoldtreatments, such that by day 14-28 scaffold was absent from the majorityof wounds. No matrix was visible by day 42.

(C) The FM Scaffold Promotes Cell Proliferation within RegeneratingWounds to a Greater Extent than Matrix Derived from Small IntestineSubmucosa

Cell proliferation within the regenerating wound can be a usefulindicator of the effectiveness of a wound treatment, as cellproliferation is indicative of a beneficial immune response, and offibroblast and/or keratinocyte migration and proliferation. In order toquantify cell proliferation, immunohistochemistry of biopsied tissueobtained during the porcine wound healing study described above wasconducted using the cell marker Ki67. Ki67 is expressed during allactive phases of the cell cycle, and is therefore a useful marker ofcell proliferation and cellular activity. Ki67 is not expressed inresting cells.

Formalin fixed biopsy tissues were mounted in paraffin and sectioned at15 μm before being subjected to citrate buffer epitope retrieval. Activecells were then detected using a Ki67 primary antibody at 1:50 dilution,and staining was developed using HRP-conjugated secondary antibody andDAB via a Bond Max™ Automated IHC/ISH Staining System (LeicaMicrosystems Instruments). Sections were then lightly counterstainedwith Mayer's haematoxylin. Immuno-stained slides were digitally imagedat 40× magnification, with three random fields taken from the epitheliallayer (days 3, 7, 14, 28 and 42), and three images taken from theregenerating dermal layer (days 7, 14, 28 and 42). Using ImageJ software(National Institute of Health), images were processed to quantify thenumber of Ki67-positive cells per frame. Firstly, images weredeconvoluted to separate the brown DAB staining from the other colorcomponents present in the image, then background was subtracted via thethreshold function. Ki67-positive cells were identified as blackclusters with a size range of 300-4500 pixels, and counted accordingly.

The quantification of cell proliferation is shown in FIG. 20, expressedas the total number of Ki67-positive cells identified in the threeepithelial frames and the three dermal frames per tissue section. Eachof the four treatment groups were sampled at the days indicated.Ki67-positive cells in the dermal layer were not quantified on day 3 dueto the absence of a dermal layer. Generally, dermal cell proliferationspiked on days 7 and 14 for all four treatment groups. On both days 7and 14, the FM scaffold treated wounds had significantly greater cellproliferation than SIS-treated and untreated groups (see FIG. 20; P<0.01one-way ANOVA using GraphPad Prism). This proliferative phase resolvedover time with Ki67-positive cells returning to ‘baseline’ on day 42 inall treatment groups.

(D) The FM Scaffold Increases Vascularization of Healing Wounds Relativeto Scaffold Derived from Small Intestine Submucosa

The presence of a functional blood supply within a wound is critical towound closure and healing. As such, increasing the vascularity of ahealing wound has received considerable attention as a means ofimproving wound healing rates and the quality of wound healing,especially in chronic wounds. The extent of vascularization (e.g.,angiogenesis) following treatment with ovine FM scaffold and smallintestine submucosa scaffold was determined in the context of theporcine wound healing model described above. In order to quantify theextent of endothelialisation and the development of vasculature in thehealing wounds, immunohistochemistry coupled with digital quantificationmethods were employed. Fixed biopsy tissues were sectioned at 15 μm,mounted and subjected to EDTA-bufferered surfactant. Endothelial cellswere stained with an anti-CD34 antibody at 1:100 dilution and visualisedusing an HRP-conjugated secondary antibody and DAB staining, beforelight counterstaining with Mayer's Haemotoxylin. Tissue biopsies takenat day 3 were excluded from this analysis given the absence of a clearlydefined dermal layer. Stained slides were digitally imaged at 40×magnification by recording four randomly selected frames of theregenerating dermal layer of each of the tissue sections. Using ImageJsoftware, the DAB color channel (CD34-positve cells) was deconvolutedfrom each of the images. The monochrome brown images were filtered toremove smaller non-specifically stained particles and smallCD34-positive clusters not representative of blood vessels (<300 μm²).The number of blood vessels was then counted using the followingcriteria; ‘small vessels’=300-500 μm², ‘medium vessels’=500-1500 μm² and‘large vessels’>1500 μm².

The average total number of vessels counted per frame is shown in FIG.21A, for each of the four treatment groups over the course of theexperiment (day 3 excluded). There was a statistically significantincrease in the number of blood vessels in wounds treated with either ofthe ovine FM scaffolds, relative to untreated wounds. The increase intotal blood vessels relative to untreated wounds was evident on days 14(P<0.01 1-ply FM and 2-ply FM), 28 (P<0.01 1-ply FM and 2-ply FM) and 42(P<0.01 1-ply FM and P<0.05 2-ply FM). In comparison, SIS treatment didnot increase the total number of blood vessels relative to the untreatedcontrol.

The number of small vessels resulting from treatment with either of theovine FM scaffolds was higher than that resulting from treatment withSIS, or from untreated groups at day 7, and this trend progressedthroughout the course of the experiment (FIG. 21C). The number ofmedium- and large-sized blood vessels was approximately equal among thefour treatment groups at day 7 of the time course (FIGS. 21D and 21E).Ovine FM-based treatments differed from SIS and untreated wounds at days14, 28 and 42 by having a greater number of medium and large vessels.For example, on day 42, ovine FM scaffold treatments had approximatelydouble the number of large vessels relative to SIS-treated and untreatedwounds (FIG. 21E).

By quantifying the numbers of small, medium and large vessels it waspossible to understand the effect of the four treatment groups on therelative size distribution of the resulting vessels. The proportions ofsmall (˜40%), medium (˜45%) and large (˜15%) vessels did not changebetween the four treatment groups over the time course (FIG. 21B). Thesefindings indicate that ovine forestomach matrix treatments increase thetotal number of vessels, but treatments do not influence the relativesize distribution of the vessels formed.

Example 18 Laminated Bioactive FM Scaffolds

Laminated FM scaffolds were developed that include an adhesive polymer,as described above. The polymer binds adjacent sheets of FM scaffold toform a laminated sheet. This polymer can also serve as a vehicle for thedelivery of bioactive molecules, and can be used to tune the release ofbioactive molecules at the site of tissue contact, as described herein.FM scaffold laminates were prepared which contained one of the followingbioactive agents in the polymer layer: growth factors FGF2 or nervegrowth factor (NGF), or antimicrobials doxycycline, amoxicillin andpoly-L-lysine. FM scaffold laminates containing the bioactive polymerlayer were subsequently assayed to demonstrate the bioactivity of thelaminate.

Specifically, a collagen gel was prepared as described in Example 3.Once formed the gel was spiked with either human recombinant FGF2 orNGF. Sterile FM scaffold sheets were cut to 8 mm discs, and growthfactor spiked collagen gel (10 μL) was applied to one surface. A secondFM scaffold disc was applied to the collagen layer to create a laminatesandwich (i.e., FM scaffold/bioactive collagen polymer/FM scaffold). Thebioactive FM scaffold laminates were dehydrated at 25° C. for 2 hours toyield a bonded laminate. The FM laminates were produced with FGF2 or NGFat either 0, 250 or 500 ng/disc. The bioactive FM scaffolds wereincubated at 37° C. for 24 hours in DMEM (1 mL) to extract the boundgrowth factor prior to application to a cell monolayer. PC12 cells wereseeded onto 24-well plates at 20 k cells/well in DMEM (1 mL) andincubated for 24 hours at 37° C. Media was removed from the cells andreplaced with DMEM (1 mL) that had been incubated with the bioactive FMscaffold laminates. Positive control cells were treated with a solutionof either FGF2 or NGF at 50 ng/mL final concentration. Cells wereincubated for 48 hours. After this time, the cell monolayer was imaged(three frames per well) using an inverted microscope. Total cells werecounted per frame, as well as the total number of cell processes perframe. Cell processes are indicative of cell differentiation asstimulated by exogenous growth factors. Cell processes were defined ascellular extensions from the cell body having a length twice that of thewidth of the cell body. Growth factor stimulated cell differentiationwas expressed as the number of cell processes per cell per frame.Results are expressed as Table 19. Both the NGF and FGF2 spikedlaminates elicited a cellular response from PC12 cells. Additionally,the response was dose-dependent, such that cellular response oflaminates spiked with 500 ng of growth factor (FGF2 or NGF) was higherthan laminates spiked with 250 ng.

TABLE 19 Quantification of the bioactivity of FGF2 and NGF containing FMscaffold laminates FGF2 Spiked Laminate (ng/disc) rhFGF2 control 500 2500 (50 ng/disc) Relative cell 0.60 ± 0.01 0.39 ± 0.02 0.09 ± 0.02 0.72 ±0.02 response NGF Spiked Laminate (ng/disc) rhNGF control 500 250 0 (50ng/disc) Relative cell 0.59 ± 0.04 0.41 ± 0.02 0.11 ± 0.02 0.73 ± 0.01response Error represent standard error from at least 6 samples

Discs of FM scaffold laminates were also prepared containing either theanti-microbial small molecule doxycycline or the polymer poly-L-lysine,as described above by spiking a collagen gel with the requiredbioactive. Doxycycline spiked discs were prepared with finalconcentrations of 30, 15, 5, 2.5, 1 and 0 μg/disc, while poly-L-lysinespiked discs were prepared a 5, 2, 1, 0.5, 0.1 and 0 μg/disc. Positivecontrols were prepared by spotting a solution of either Doxycycline(final concentration 30 μg/disc) or poly-L-lysine (final concentration 5μg/disc) onto sterile filter paper discs. Plates of Mueller-Hinton agarwere prepared and streaked with 100 μL of 10⁸ S. aureus and left to dryfor 5-10 minutes. Using sterile technique, discs were transferred to theplates and the plates incubated for 24 hours at 35° C. After 24 hours,plates were digitally imaged and the anti-microbial zone of inhibitionaround each of the discs was scored relative to the positive control.Bioactivity against S. aureus of the laminated FM scaffolds is given inTable 20. Bioactivity of the test articles were scored, where ‘+++’indicates a bioactivity comparable to the positive control, and ‘−’indicates no bioactivity.

TABLE 20 Bioactivity against S. aureus of FM scaffolds laminated witheither Doxycycline or poly-L-lysine Positive Antibiotic concentrationcontrol Doxycycline μg/disc 30 15 5 2.5 1 0 30 Antibacterial +++ +++++ + − − +++ score Poly-L- lysine μg/disc 5 2 1 0.5 0.1 0 5Antibacterial +++ − − − − − +++ score

Discs of FM scaffold previously laminated with a collagen polymercontaining doxycycline were bioactive against cultures of S. aureus.Bioactivity of the doxycycline laminated FM scaffolds were concentrationdependant such that laminated discs at 30 μg/disc had approximatelyequivalent bioactivity to the control discs, while laminates at 1 and 0μg/disc demonstrated no bioactivity. Bioactivity of FM scaffold laminatecontaining poly-L-lysine was less pronounced than doxycycline FMscaffold laminates. However, bioactivity was observed at the highestconcentration tested of poly-L-lysine (5 pg/disc), which compared wellwith the control disc.

Results of these in vitro studies demonstrated the ability of growthfactor proteins or anti-microbial small molecules and polymers todiffuse with time from the polymeric adhesive layer. Thus, the resultsindicate that a laminated FM sheet which contains a polymer layercomprising a bioactive agent may be used for a variety of treatmentpurposes. The laminate can range from 2-ply to 15-ply or more (e.g., 2to 30-ply) depending on specific clinical applications and hence therequired extent of physical performance.

In vivo, the bioactive FM scaffold laminate similarly diffuses bioactiveagent from the polymer layer to surrounding tissues. By varying the typeof polymer layer and the formulation of the bioactive agent within thislayer, the diffusion of bioactive agent from the laminate to localtissue can be controlled.

Example 19 Biocompatibility of Ovine FM Scaffolds

Ovine FM scaffolds were prepared according to Example 1, cut to 4×4 cmsquare devices and terminally sterilized using ethylene oxide. FMscaffold devices were tested for biocompatibility according to Blue BookMemorandum G95-1 and ISO10993-1:2003. Testing included cytotoxicity,sensitization and irritation/intracutaneous reactivity. In an effort todetect any microbial infection resulting from the FM scaffold device,and/or detect any immunological or inflammatory response, the requiredtesting was expanded to include a 30-day in vivo implantation assay.Endotoxin concentrations of the devices were also quantified.

All devices and controls were prepared according to ISO10993-12:2007‘Biological Evaluation of Medical Devices, Part 12: Sample Preparationand Reference Materials’. The results of biocompatibility testing aresummarized in Table 21, and indicate that the biocompatibilitycharacteristics of FM scaffolds make them suitable for in vivo delivery.

TABLE 21 Biocompatibility testing of FM scaffold Biocompatibility testTest description Result Cytotoxicity Cytotoxicity Test Using the ISONon-cytotoxic Elution Method in a L-929 Mouse Fibroblast Cell Lineaccording to ISO 10993-5:1999 ‘Biological Evaluation of Medical Devices,Part 5: Tests for In Vitro Cytotoxicity’ Cytotoxicity Agar Overlayaccording to ISO Non-cytotoxic 10993-5:1999 ‘Biological Evaluation ofMedical Devices, Part 5: Tests for In Vitro Cytotoxicity’ Irritation ISO10993-10:2002, Amendment Non-irritating 1:2006, ‘Biological Evaluationof Medical Devices, Part 10; Tests for Irritation and Delayed-TypeHypersensitivity’ Sensitization According to ISO 10993-10:2002,Non-sensitizing Amendment 1:2006, ‘Biological Evaluation of MedicalDevices, Part 10; Tests for Irritation and Delayed- TypeHypersensitivity’ 30 day According to ISO10993-6:1994 Non-irritatingImplantation ‘Biological Evaluation of Medical Devices, Part 6: Testsfor Local Effects After Implantation’ Endotoxin USP<85>, ANSI/AAMIST72:2002 Passed Testing and FDA guidelines

Example 20 Viral Inactivation During Manufacture of Ovine FM Scaffolds

Viral inactivation during the production of FM scaffold can occur as aresult of: 1) viral protein denaturation and/or disruption of virallipids of the viral envelope during treatment with detergents (TritonX-200 and SDS); 2) the liberation of reactive oxygen species duringperacetic acid (PAA) treatment; and 3) terminal sterilization usingethylene oxide. A study was undertaken to examine viral inactivationduring the STOF processes and PAA treatment using three model viruses.

The level of viral inactivation was tested using a scaled-downmanufacturing process. This involved spiking high-titre viral stocks ofeach of the three model viruses into samples representing two differentstages of the manufacturing process, and subjecting these spiked samplesto a treatment analogous to the corresponding manufacturing step. Thelevel of virus inactivation during each treatment was determined bycomparing the viral titres recovered from the spiked samples after eachtreatment, with viral recovery from untreated samples.

The model viruses were chosen to include viruses with differentphysio-chemical characteristics, such as the presence or absence ofenvelope, type of nucleic acid genome (DNA or RNA) and survival in theenvironment/resistance to disinfection (Table 22). The amount ofinfectious virus present in samples was determined by end-pointtitration and viral titres were expressed as tissue culture infectivedose, 50% (TCID₅₀).

The model systems employed in this study demonstrated that both STOF andPAA treatments inactivated the model viruses, with a total theoreticalreduction in viral titer of more than 6 logs TCID₅₀ (Table 22).

The detergent treatment was effective at inactivating enveloped viruses(PI-3 and FHV-1), but showed little effect at inactivating thenon-enveloped virus BEV. This result was expected, as non-envelopedviruses are typically not affected by treatment with detergents. Incontrast, PAA treatment was effective at inactivating all three modelviruses (Table 22).

TABLE 22 Demonstrated and theoretical inactivation of three modelviruses during the scaled-down manufacturing process of FM scaffoldTotal STOF PAA** theoretical inactivation Inactivation inactivationNatural (log (log (log Virus (family) host Genome Envelope ResistanceTCID₅₀)* TCID₅₀) TCID₅₀) PI-3 Bovine ss(−)RNA Yes Sensitive =4.0* =2.3=6.3 FHV-1 Feline ds DNA Yes Sensitive =3.5 =2.6 =6.1 BEV Boviness(+)RNA No Resistant 0.5 7.3 7.3 Abbreviations used: PI-3:Parainfluenza virus type 3 (Paramyxoviridae); FHV-1: Felid herpesvirustype 1 (Herpesviridae); BEV: Bovine enterovirus (Picornaviridae);TCID50: Tissue culture infective dose, 50%, PAA: peracetic acid.*Inactivation of model viruses following treatment with two detergentsolutions in the presence of an osmotic gradient. Detergent solutionsincluded 0.1% EDTA/0.028% Triton X-200 and 0.1% SDS. **Inactivation ofmodel viruses following treatment with 0.3% PAA/5% Ethanol in PBS. Theresults are expressed as “more or equal to” when the lower limit ofvirus recovery was limited by toxicity of the samples to mammaliancells.

Example 21 Fabrication of a Device for Breast Reconstruction

Wet sheets of naturally concave FM scaffold were aligned andsuccessively layered onto one another to create 4-ply, 6-ply, and 8-plylaminates. The sheets were layered onto one another using a curved formthat supported and complemented the natural contours of the FM scaffold.The laminates were freeze-dried to yield bonded laminates. Theselaminates were subsequently sewn together using a straight-stitch ofvicryl suture, with a stitch length of approximately 3 mm. Laminateswere sewn in straight lines following the curvature of the laminate.Stitch lines were separated by approximately 10 mm. The breast laminateswere of either a crescent or ellipse shape, as illustrated in FIGS. 2Aand 2B, respectively. The breast laminates can be affixed to nativetissue using sutures or staples.

REFERENCES

-   Boguszewski, D. V., N. A. Dyment, et al. (2008). Biomechanical    Comparison of Abdominal Wall Hernia Repair Materials. ASME 2008    Summer Bioengineering Conference Marriott Resort, Marco Island,    Fla., ASME-   Choe, J. M., R. Kothandapani, et al. (2001). “Autologous, cadaveric,    and synthetic materials used in sling surgery: comparative    biomechanical analysis.” Urology 58(3): 482-6.-   Freytes, D. O., R. S. Tullius, et al. (2006). “Effect of storage    upon material properties of lyophilized porcine extracellular matrix    derived from the urinary bladder.” J Biomed Mater Res B Appl    Biomater 78(2): 327-33.-   Freytes, D. O., R. S. Tullius, et al. (2008). “Hydrated versus    lyophilized forms of porcine extracellular matrix derived from the    urinary bladder.” J Biomed Mater Res A 87(4): 862-72.-   Gouk, S. S., T. M. Lim, et al. (2008). “Alterations of human    acellular tissue matrix by gamma irradiation: histology,    biomechanical property, stability, in vitro cell repopulation, and    remodeling.” J Biomed Mater Res B Appl Biomater 84(1): 205-17.-   Lemer, M. L., D. C. Chaikin, et al. (1999). “Tissue strength    analysis of autologous and cadaveric allografts for the pubovaginal    sling.” Neurourol Urodyn 18(5): 497-503.-   Lindblad, W. J. (2006). “How should one study wound healing?” Wound    Repair Regen 14(5): 515.-   Lindblad, W. J. (2008). “Considerations for selecting the correct    animal model for dermal wound-healing studies.” J Biomater Sci Polym    Ed 19(8): 1087-96.-   Morgan, A. S., T. McIff, et al. (2004). “Biomechanical properties of    materials used in static facial suspension.” Arch Facial Plast Surg    6(5): 308-10.-   Sclafani, A. P., S. A. McCormick, et al. (2002). “Biophysical and    microscopic analysis of homologous dermal and fascial materials for    facial aesthetic and reconstructive uses.” Arch Facial Plast Sure    4(3): 164-71.-   Vural, E., N. McLaughlin, et al. (2006). “Comparison of    biomechanical properties of alloderm and enduragen as static facial    sling biomaterials.” Laryngoscope 116(3): 394-6.-   Zerris, V. A., K. S. James, et al. (2007). “Repair of the dura mater    with processed collagen devices.” J Biomed Mater Res B Appl Biomater    83(2): 580-8.

EQUIVALENTS

The invention has been described herein with reference to certainexamples and embodiments only. No effort has been made to exhaustivelydescribe all possible examples and embodiments of the invention. Indeed,those of skill in the art will appreciate that various additions,deletions, modifications and other changes may be made to theabove-described examples and embodiments, without departing from theintended spirit and scope of the invention as recited in the followingclaims. It is intended that all such additions, deletions, modificationsand other changes be included within the scope of the following claims.

INCORPORATION BY REFERENCE

The entire contents of all patents, published patent applications,websites, and other references cited herein are hereby expresslyincorporated herein in their entireties by reference.

1. A tissue scaffold comprising the extracellular matrix of thepropria-submucosa of the forestomach of a ruminant.
 2. The tissuescaffold of claim 1, wherein the propria-submucosa is from the rumen. 3.The tissue scaffold of claim 1, further comprising decellularised tissueselected from the group consisting of epithelium, basement membrane,tunica muscularis, and combinations thereof.
 4. The tissue scaffold ofclaim 1, further comprising a fibrillar protein selected from the groupconsisting of collagen I, collagen III, elastin, and combinationsthereof.
 5. The tissue scaffold of claim 1, further comprising a growthfactor selected from the group consisting of FGF-2, TGFb1, TGFb2, VEGF,and combinations thereof.
 6. The tissue scaffold of claim 1, furthercomprising a glycosaminoglycan selected from the group consisting ofhyaluronic acid, heparan sulfate, and combinations thereof.
 7. Thetissue scaffold of claim 1, further comprising an adhesive proteinselected from the group consisting of fibronectin, laminin, collagen IV,and combinations thereof.
 8. The tissue scaffold of claim 1, having acontoured luminal surface.
 9. The tissue scaffold of claim 1, formattedas a single or laminated sheet.
 10. The tissue scaffold of claim 9,comprising 2-12 laminated sheets.
 11. The tissue scaffold of claim 10,further comprising a polymer positioned between two or more laminatedsheets.
 12. The tissue scaffold of claim 11, wherein the polymer isselected from the group consisting of collagen, chitosan, alginate,polyvinyl alcohol, carboxymethyl cellulose, hydroxypropyl cellulose, andcombinations thereof.
 13. The tissue scaffold of claim 11, wherein thepolymer further comprises a bioactive molecule.
 14. The tissue scaffoldof claim 13, wherein the bioactive molecule is dispersed throughout alayer of the polymer.
 15. The tissue scaffold of claim 13, wherein thebioactive molecule is non-covalently linked to the polymer.
 16. Thetissue scaffold of claim 13, wherein the bioactive molecule iscovalently linked to the polymer.
 17. The tissue scaffold of claim 13,wherein the bioactive molecule is a small molecule or a polypeptide. 18.The tissue scaffold of claim 17, wherein the bioactive molecule isselected from the group consisting of a growth factor, ananti-microbial, an analgesic, a hemostatic, a pro-angiogenic agent, ananti-angiogenic agent, and combinations thereof.
 19. The tissue scaffoldof claim 18, wherein the bioactive molecule is selected from the groupconsisting of FGF2, NGF, doxycycline, poly-L-lysine, and combinationsthereof.
 20. The tissue scaffold of claim 10, wherein the laminatedsheets are secured together by stitches or sutures.
 21. The tissuescaffold of claim 1, wherein the sheet has a width of at least 10 cm anda length of at least 10 cm.
 22. The tissue scaffold of claim 1, whereinthe sheet has an average burst strength of at least 80N.
 23. The tissuescaffold of claim 1, wherein the ruminant belongs to a genus selectedfrom the group consisting of Capra, Bos, Cervus and Ovis.
 24. The tissuescaffold of claim 1, wherein the ruminant is Ovis aries.
 25. Acomposition comprising the tissue scaffold of claim
 1. 26. Use of thetissue scaffold of claim 1 to cover a tissue deficit.
 27. The use ofclaim 26, wherein the tissue deficit has a width of at least 10 cm and alength of at least 10 cm.
 28. The use of claim 26, wherein the tissuescaffold increases proliferation of cells within the tissue deficit. 29.The use of claim 26, wherein the tissue scaffold increasesvascularization within the tissue deficit.
 30. Use of the tissuescaffold of claim 1 to reinforce soft tissue.
 31. The use of claim 30,wherein the reinforced soft tissue is repaired with a scaffold having awidth of at least 10 cm and a length of at least 10 cm.
 32. A method forinducing repair of a damaged tissue, comprising contacting the damagedtissue with a tissue scaffold of claim
 1. 33. A method for stimulatingsoft tissue regeneration, comprising contacting the soft tissue with atissue scaffold of claim
 1. 34. The method of claim 32 or 33, whereinthe tissue scaffold induces cell proliferation.
 35. The method of claim32 or 33, wherein the tissue scaffold induces vascularization.
 36. Amethod of inducing vascularization of a tissue, comprising contactingthe tissue with a tissue scaffold of claim 1, such that vascularizationoccurs within the tissue.
 37. A method of separating or decellularisingthe layers within all or a portion of a tissue, comprising creating atransmural osmotic flow between two sides of the tissue, such that thelayers within all or a portion of the tissue are separated ordecellularised.
 38. The method of claim 37, wherein transmural osmoticflow is created from the luminal to the abluminal side of the tissue.39. The method of claims 37, comprising removing all or part of a tissuelayer selected from the group consisting of epithelium, basementmembrane, tunica muscularis, and combinations thereof.
 40. The method ofclaim 37, wherein the tissue comprises a keratinized stratified squamousepithelium.
 41. The method of claim 37, further comprising the step ofencapsulating a first solution within the tissue or portion thereof, andimmersing the tissue or portion thereof in a second solution which ishypertonic to the first solution.
 42. The method of claim 41, furthercomprising the step of removing the tissue or portion thereof from thesecond solution, and immersing the tissue or portion thereof in a thirdsolution which is also hypertonic to the first solution.
 43. The methodof claim 37, further comprising the step of encapsulating a firstsolution within the tissue or portion thereof, and immersing the tissueor portion thereof in a second solution which is hypotonic to the firstsolution.
 44. The method of claim 43, further comprising the step ofremoving the tissue or portion thereof from the second solution, andimmersing the tissue or portion thereof in a third solution which isalso hypotonic to the first solution.
 45. The method of claim 43,wherein the second solution comprises water, optionally including atleast one buffer, detergent or salt.
 46. The method of claim 43, whereinthe first solution comprises 4 M NaCl.
 47. The method of claim 44,wherein the first solution comprises 4 M NaCl, the second solutioncomprises 0.028% Triton X-200 and 0.1% EDTA, and the third solutioncomprises 0.028% SDS.
 48. The method of claim 44, wherein the firstsolution comprises 4 M NaCl, the second solution comprises 0.1% SDS, andthe third solution comprises 0.028% Triton X-200 and 0.1% EDTA.
 49. Themethod of claim 37, wherein the method is performed at a temperaturebetween 2° C. and 4° C. in less than 36 hours.
 50. The method of claim37, wherein the method is performed at 4° C.
 51. The method of claim 37,wherein the method is performed in less than 24 hours.
 52. The method ofclaim 46, wherein the second solution comprises 0.028% Triton X-200,0.1% EDTA, and 0.1% SDS.
 53. The method of claim 52, wherein the methodis performed at a temperature between 18° C. and 24° C.
 54. The methodof claims 52, wherein the method is performed in less than 6 hours. 55.The method of claim 37, wherein the tissue or portion thereof isdistended to increase the transmural osmotic flow.
 56. The method ofclaim 37, wherein the tissue is a whole organ, or a portion thereof. 57.The method of claim 37, wherein the tissue is derived from theforestomach of a ruminant.
 58. The method of claim 57, wherein thetissue is the rumen.
 59. The method of claim 57, wherein the ruminantbelongs to a genus selected from the group consisting of Capra, Bos,Cervus and Ovis.
 60. The method of claim 59, wherein the ruminant isOvis aries.
 61. A tissue scaffold produced according to the method ofclaim
 37. 62. A method of forming a tissue scaffold, comprisingperforming the method of claim
 37. 63. An implantable tissue scaffolddevice for supporting breast tissue within a patient, wherein saiddevice comprises extracellular matrix of the propria-submucosa of theforestomach of a ruminant.
 64. The device of claim 63, wherein thebreast tissue comprises a prosthesis.
 65. The device of claim 63,formatted as a laminated sheet comprising two or more layers ofextracellular matrix.
 66. The device of claim 65, wherein the laminatedsheet comprises 4-8 layers of extracellular matrix.
 67. The device ofclaim 63, which has concavity.
 68. The device of claim 65, wherein thelayers of extracellular matrix are secured together by stitches orsutures.
 69. The device of claim 63, wherein the extracellular matrix isperforated.
 70. The device of claim 63, wherein the extracellular matrixis unperforated.
 71. The device of claim 63, wherein the device has acrescent shape.
 72. The device of claim 63, wherein the device has anelliptical shape.
 73. A method of supporting breast tissue within apatient, comprising positioning the device of claim 63 within thepatient in a supporting position relative to the breast tissue.
 74. Themethod of claim 73, wherein the breast tissue comprises a breastprosthesis.
 75. The method of claim 73, wherein the breast tissuecomprises native tissue.
 76. The method of claim 73, wherein saidpositioning comprises covering the lower and lateral sections of thebreast tissue.
 77. A tissue scaffold comprising two or more sheets ofextracellular matrix, laminated by a polymer positioned between thesheets.
 78. The tissue scaffold of claim 77, wherein the scaffoldcomprises extracellular matrix of the submucosa of a tissue selectedfrom the group consisting of small intestine, stomach, bladder,pericardium and dermis.
 79. The tissue scaffold of claim 77, wherein theextracellular matrix comprises collagen.
 80. The tissue scaffold ofclaim 77, wherein the polymer is selected from the group consisting ofcollagen, chitosan, alginate, polyvinyl alcohol, carboxymethylcellulose, hydroxypropyl cellulose, and combinations thereof.
 81. Thetissue scaffold of claim 77, wherein the polymer further comprises abioactive molecule.
 82. The tissue scaffold of claim 81, wherein thebioactive molecule is non-covalently linked to the polymer.
 83. Thetissue scaffold of claim 81, wherein the bioactive molecule iscovalently linked to the polymer.
 84. The tissue scaffold of claim 81,wherein the bioactive molecule is a small molecule or a polypeptide. 85.The tissue scaffold of claim 84, wherein the bioactive molecule isselected from the group consisting of a growth factor, ananti-microbial, an analgesic, a hemostatic, a pro-angiogenic agent, ananti-angiogenic agent, and combinations thereof.
 86. The tissue scaffoldof claim 85, wherein the bioactive molecule is selected from the groupconsisting of FGF2, NGF, doxycycline, poly-L-lysine, and combinationsthereof.